key: cord-1041668-fcalycpw
authors: Metzemaekers, Mieke; Cambier, Seppe; Blanter, Marfa; Vandooren, Jennifer; de Carvalho, Ana Carolina; Malengier‐Devlies, Bert; Vanderbeke, Lore; Jacobs, Cato; Coenen, Sofie; Martens, Erik; Pörtner, Noëmie; Vanbrabant, Lotte; Van Mol, Pierre; Van Herck, Yannick; Van Aerde, Nathalie; Hermans, Greet; Gunst, Jan; Borin, Alexandre; Toledo N Pereira, Bruna; dos SP Gomes, Arilson Bernardo; Primon Muraro, Stéfanie; Fabiano de Souza, Gabriela; S Farias, Alessandro; Proenca‐Modena, José Luiz; R Vinolo, Marco Aurélio; Marques, Pedro Elias; Wouters, Carine; Wauters, Els; Struyf, Sofie; Matthys, Patrick; Opdenakker, Ghislain; Marques, Rafael Elias; Wauters, Joost; Gouwy, Mieke; Proost, Paul
title: Kinetics of peripheral blood neutrophils in severe coronavirus disease 2019
date: 2021-04-29
journal: Clin Transl Immunology
DOI: 10.1002/cti2.1271
sha: 216ba57657f06023f5c10c120c3d1c2f00c38efc
doc_id: 1041668
cord_uid: fcalycpw

OBJECTIVES: Emerging evidence of dysregulation of the myeloid cell compartment urges investigations on neutrophil characteristics in coronavirus disease 2019 (COVID‐19). We isolated neutrophils from the blood of COVID‐19 patients receiving general ward care and from patients hospitalised at intensive care units (ICUs) to explore the kinetics of circulating neutrophils and factors important for neutrophil migration and activation. METHODS: Multicolour flow cytometry was exploited for the analysis of neutrophil differentiation and activation markers. Multiplex and ELISA technologies were used for the quantification of protease, protease inhibitor, chemokine and cytokine concentrations in plasma. Neutrophil polarisation responses were evaluated microscopically. Gelatinolytic and metalloproteinase activity in plasma was determined using a fluorogenic substrate. Co‐culturing healthy donor neutrophils with severe acute respiratory syndrome coronavirus 2 (SARS‐CoV‐2) allowed us to investigate viral replication in neutrophils. RESULTS: Upon ICU admission, patients displayed high plasma concentrations of granulocyte–colony‐stimulating factor (G‐CSF) and the chemokine CXCL8, accompanied by emergency myelopoiesis as illustrated by high levels of circulating CD10(−), immature neutrophils with reduced CXCR2 and C5aR expression. Neutrophil elastase and non‐metalloproteinase‐derived gelatinolytic activity were increased in plasma from ICU patients. Significantly higher levels of circulating tissue inhibitor of metalloproteinase 1 (TIMP‐1) in patients at ICU admission yielded decreased total MMP proteolytic activity in blood. COVID‐19 neutrophils were hyper‐responsive to CXCL8 and CXCL12 in shape change assays. Finally, SARS‐CoV‐2 failed to replicate inside human neutrophils. CONCLUSION: Our study provides detailed insights into the kinetics of neutrophil phenotype and function in severe COVID‐19 patients, and supports the concept of an increased neutrophil activation state in the circulation.

The pandemic of COVID-19 and SARS-CoV-2, first identified in Wuhan (China) in December 2019, has resulted in over 115 million cases and more than 2.5 million deaths (WHO, covid19.who.int, as of March 2021). SARS-CoV-2 is a novel coronavirus of zoonotic origin that uses angiotensinconverting enzyme II (ACE2) as main receptor. 1 The majority of patients remain either asymptomatic or develop mild clinical symptoms such as fever, shortness of breath, cough, myalgia and fatigue. [2] [3] [4] However, 10-20% of the patients develop severe viral pneumonia and acute respiratory distress syndrome (ARDS), associated with respiratory failure. 2, 3 Often, admission to intensive care units (ICUs) and mechanical ventilation are needed, and a considerable proportion of patients evolve to multiple-organ failure and eventually succumb. 5 Patients suffering from severe COVID-19 frequently have increased coagulopathy and may develop thrombotic events and neurological symptoms. 6, 7 Common comorbidities such as hypertension, diabetes and asthma, as well as older age, are considered risk factors for severe and potentially fatal COVID-19. 4, 8 Severe COVID-19 cases present with a systemic inflammatory response accompanied by elevated levels of acute-phase reactants, cytokines and chemokines [e.g. C-reactive protein (CRP), soluble IL-2 receptor (IL-2R), IL-6, CXCL8, CCL2, CCL5, IL-10 and tumor necrosis factor (TNF)-a], the latter being referred to as cytokine storm. 4, [9] [10] [11] White blood cell analysis revealed that a dysregulated host immune response features severe COVID-19. Abnormally low lymphocyte (especially Tlymphocytes) counts, as well as lower percentages of monocytes, basophils and eosinophils, have been described in (severe) COVID-19 cases. 4, 7, 9, 10, 12 Moreover, severe cases have higher neutrophil counts and a higher neutrophil-to-lymphocyte ratio (NLR), 4,5 which appeared as an independent biomarker predicting critical illness and poor prognosis. [13] [14] [15] Neutrophils are highly versatile granulocytes constituting 50-70% of all peripheral blood leucocytes in humans and are the first responders to infection and tissue damage. 16 After recruitment to sites of infection and inflammation by the local action of chemokines (e.g. CXCL8), neutrophils can phagocytose and kill pathogens, release cytotoxic compounds and degradative enzymes, produce reactive oxygen species (ROS) and expel neutrophil extracellular traps (NETs) [17] [18] [19] Moreover, phenotypically distinct neutrophil subpopulations or divergently polarised cells with functional diversity were recently identified under homeostatic and pathological conditions. 20, 21 Inappropriate neutrophil activation (hyperactivation) can lead to excessive inflammation and collateral injury to (healthy) tissue as seen in subjects with several (lung) inflammatory diseases. 21, 22 Given the fact that circulating neutrophils are increased both in absolute numbers and in percentage in patients with severe COVID-19, we collected blood samples from COVID-19 patients hospitalised in dedicated general wards or in ICUs, and characterised neutrophils at phenotypical and functional levels during disease evolution. We compared the parameters with those from healthy volunteers.

Cytokine/chemokine levels were quantified in plasma from healthy controls and COVID-19 patients during the disease course. Plasma G-CSF levels were significantly higher in patients hospitalised in general wards and at ICU days 1 and 7 as compared to levels detected in healthy donors ( Figure 1a ). Upon discharge from the ICU, G-CSF plasma concentrations from most ICU patients were decreased to levels resembling those detected in healthy controls. Circulating granulocyte-macrophage colony-stimulating factor (GM-CSF) concentrations were < 1 pg mL À1 for most donors, and no differences between the study groups were detected (data not shown). The levels of CXCL12a, known to retain immature neutrophils in the bone marrow microenvironment, 23 peaked in the bloodstream after 1 week at ICU (Figure 1b) . A tendency for increased CXCL12a was observed for other patient groups (Figure 1b) . In contrast to the unaltered levels of the neutrophil chemoattractants CXCL1 and CXCL5, the most powerful human neutrophilattracting chemokine CXCL8 was significantly more abundant in plasma from ICU and ward patients as compared to healthy donors (Figure 1c-e) . Similar to G-CSF concentrations, CXCL8 levels significantly decreased towards ICU discharge. Compared to healthy controls, COVID-19 patients showed significantly higher plasma levels of the highly potent CXCR3 ligands CXCL10 and CXCL11 (Figure 1f, g) . Moreover, the activity in plasma of CD26/dipeptidyl peptidase IV (DPP4), a highly specific serine protease that inactivates CXCL10, CXCL11 and CXCL12 within minutes, 24 was lower in all patient groups than in healthy controls ( Figure 1h) . Altogether, these results show the excessive release of neutrophilactivating and neutrophil-mobilising factors during severe COVID-19, setting the stage for a scenario of neutrophil hyperactivation.

In order to evaluate the responsiveness of neutrophils from COVID-19 patients, a microscopic evaluation of blood neutrophil polarisation was performed in the presence or absence of proinflammatory mediators. As expected, few neutrophils from COVID-19 patients and healthy controls were polarised directly after isolation (baseline) or upon exposure to vehicle (Figure 2a , b). Remarkably, at ICU day 7 (but not at ICU admission), neutrophils were hyper-responsive to CXCL8 and CXCL12a in comparison with control cells (Figure 2c -e). No significant polarisation was elicited with TNF-a or CXCL10 (Figure 2f-h) . No significant differences in polarisation responses were seen if neutrophils from healthy donors were exposed to plasma from COVID-19 patients versus plasma from healthy subjects (Figure 2i ).

Neutrophils are packed with readily mobilisable granules containing pre-made inflammatory mediators. Among these pre-synthesised effector molecules are the serine protease neutrophil elastase and neutrophil gelatinase or matrix metalloproteinase-9 (MMP-9). We monitored the plasma levels of these enzymes to gain insight into the activation state of circulating neutrophils (Table 1) . Neutrophil elastase concentrations were significantly enhanced in plasma from ICU patients as compared to healthy volunteers, indicating peripheral neutrophil activation in these patients (Figure 3a ). The highest neutrophil elastase levels were detected upon ICU admission (Figure 3a ). These observations are in line with the previously reported enhanced myeloperoxidase (MPO) levels in COVID-19 plasma. 25 A trend towards enhanced neutrophil CXCL8, (f) CXCL10 and (g) CXCL11 in plasma samples from COVID-19 patients who stayed at the ICU [samples were collected during the first 48 h after admission (ICUday 1; n ≥ 10), after one week (ICUday 7; n ≥ 10) and upon discharge from the ICU (ICUdischarge; n ≥ 10)], COVID-19 patients who were hospitalised in general wards (ward; n ≥ 13) and healthy controls (HC; n ≥ 7). Moreover, (h) CD26/DPP4 enzyme activity was determined in a substrate conversion assay. Bars indicate the median plasma cytokine/chemokine concentration (a-g) or CD26 activity (h) for each study group. Results were statistically analysed by the Kruskal-Wallis with Dunn's multiple comparisons tests. *P ≤ 0.05; **P ≤ 0.01; ***P ≤ 0.001; ****P ≤ 0.0001 for statistical differences between patients and controls. $P ≤ 0.05; $$$P ≤ 0.001; $$$$P ≤ 0.0001 for statistical differences between ICUday 1 and other patient groups. Polarisation responses of peripheral blood neutrophils from patients with COVID-19. Neutrophils and plasma were collected from the peripheral blood of COVID-19 patients who stayed at the ICU [samples were collected during the first 48 h after admission (ICUday 1; n ≥ 7), after one week (ICUday 7; n ≥ 5) and upon discharge from the ICU (ICUdischarge; n ≥ 7)], COVID-19 patients who were hospitalised in general wards (ward; n = 9) and healthy controls (HC; n = 6). Neutrophils were resuspended in HBSS buffer, incubated in the presence of a chemotactic stimulus and fixed, upon which the percentage of polarised cells (as determined by the cellular shape) was determined microscopically. (a) Baseline polarisation was determined by fixing the cells immediately after purification. Stimulus-induced polarisation was determined by incubating the cells for 3 min in the presence of the following stimuli: (b) vehicle; (c) CXCL8 (5 ng mL À1 ); (d) CXCL8 (12.5 ng mL À1 ); (e) CXCL12 (300 ng mL À1 ); (f) TNF-a (50 ng mL À1 ); (g) CXCL10 (100 ng mL À1 ); and (h) CXCL10 (300 ng mL À1 ). Moreover, (i) healthy donor neutrophils were treated with plasma from COVID-19 patients or healthy donors. Results are represented as percentage of polarised cells and were statistically analysed by the Kruskal-Wallis with Dunn's multiple comparisons tests. *P ≤ 0.05 for statistical differences between groups indicated by horizontal lines. elastase levels was found in plasma from ward patients ( Figure 3a) . Interestingly, as compared to healthy controls, the level of circulating TIMP-1 (an endogenous metalloproteinase inhibitor) was significantly higher in ICU patients at the time of admission, and a tendency towards elevated TIMP-1 concentrations was detected in all other patient groups (Figure 3b ). TIMP-1/MMP-9 complexes were significantly more abundant in plasma from ICU patients (Figure 3c ). Plasma concentrations of the pan proteinase inhibitor alpha-2macroglobulin (a2M) were found highly variable within the different study groups. Although no significant differences were detected between groups, some patients presented with three-to 10-fold higher a2M concentrations than healthy controls (Figure 3d ).

Given that neutrophil elastase and MMP-9 are proteases with gelatinolytic activity, we measured the gelatinolytic activity in patient plasma (Table 1) . Although no significant differences were found in total gelatinolytic activity, nonmetalloproteinase-derived gelatinolytic activity was significantly increased in plasma from COVID-19 patients seven days after admission (Figure 3e , f). Interestingly, total MMP activity was significantly decreased in plasma from COVID-19 patients upon admission to ICU and general wards (Figure 3g , h). In conclusion, the products of activated neutrophils and nonmetalloproteinase-derived gelatinolytic activity are increased in the blood from patients with severe COVID-19, whereas increased TIMP-1 levels may contribute to counterbalancing the increases in MMP-9.

Evidence for immature, activated neutrophils in peripheral blood from patients with severe COVID-19

Peripheral blood neutrophils were examined for the expression of adhesion molecules, activation/maturation markers and chemoattractant receptors using multicolour flow cytometry. Phenotypical characterisation of neutrophils revealed the presence of immature, activated cells in peripheral blood from COVID-19 patients (Figure 4 ). Upon ICU admission, 45-90% of blood neutrophils were immature as evidenced by the lack of CD10 expression and the reduced levels of the low-affinity FccR CD16 26, 27 (Figure 4a, b) . These observations indicate a situation of emergency myelopoiesis. 16, 28 In the course of their ICU stay, COVID-19 patients gradually acquired a higher percentage of mature neutrophils in the circulation, as the CD10 + population increased (Figure 4a ). However, for the vast majority of patients, the relative abundance of mature neutrophils was still lower than that of healthy controls (Figure 4a) . Plasma levels of G-CSF correlated inversely with the percentage of mature neutrophils (Supplementary figure 1) .

During the ICU stay, multiple signs of neutrophil activation were observed. The chemokine receptor CXCR2, but not CXCR1, was significantly less abundant on patient neutrophils at ICU day 1 and day 7 as compared to controls (Figure 4c, d) . As expected, most patients did not present any alterations in the expression levels of the chemokine receptors CXCR3 and CXCR4 during infection (Supplementary figure 2a and b) . Expression of complement receptor C5aR was significantly lower upon ICU admission as compared to healthy donors ( Figure 4e ). Moreover, increased CD66b levels were detected at the beginning of the ICU stay versus discharge (Figure 4f ). Complement receptor 1 (CR1) or CD35 was significantly more abundant on neutrophils from ICU patients at day 7 of hospitalisation as compared to admission (Figure 4g ). The expression of the tetraspanin CD63, a surface receptor for TIMP-1, 29 was increased after one week of hospitalisation at ICU as compared to discharge (Figure 4h ). No expression of CD49d, IL-1R1, CCR1, CCR2 or ICAM-1 was detected in any group. For IL-1R2, BLTR1, CD11b, CD11c, CD15, CD62L, FPR1 and HLA-DR, no significant differences were found between patients and controls, although a tendency towards increased numbers of HLA-DR positive cells was seen in the circulation from COVID-19 patients (Supplementary figure 2c-j) .

A pertinent question in the case of COVID-19 is whether neutrophils may propagate the virus. Indeed, in view of the phenotypical and functional alterations of peripheral blood neutrophils from COVID-19 patients, it was imperative to investigate possible diseasepromoting effects. To explore whether SARS-CoV-2 is capable of infecting neutrophils, we incubated freshly isolated neutrophils from healthy donors with SARS-CoV-2 or vehicle in vitro. SARS-CoV-2 infectivity and ability to replicate in cells were validated in permissive Vero CCL81 cells (Figure 5a,  b) . Evaluation of the viral load in neutrophils 6 or 12 h post-infection revealed low static levels of viral RNA inside cells and in culture supernatant, and decreasing levels of infective SARS-CoV-2 in culture supernatant, indicating that SARS-CoV-2 is unable to replicate in human neutrophils in vitro (Figure 5c-e) . In addition, microscopic evaluation of neutrophil cell cultures infected with SARS-CoV-2 confirmed the lack of virus-induced cytopathic effects or cell death (Figure 5f ). Accordingly, the levels of lactate dehydrogenase (LDH)that increase upon neutrophil deathin supernatant were not increased if cells were infected with SARS-CoV-2 as compared to vehicle-treated cells (Figure 5g ). The assessment of MPO activity revealed similar enzymatic activities in supernatants from SARS-CoV-2-challenged and vehicle-treated neutrophils (Figure 5h ). An increase in MPO release was observed from 6 to 12 h postinfection, but was similar between the virus-and vehicle-treated groups.

The COVID-19 pandemic has come on top of the annual death toll of infectious diseases and remains a significant risk to human health worldwide. The precise mechanisms underlying the heterogeneity of COVID-19 disease courses are incompletely understood, although an advanced age and/or the presence of comorbidities that provide a certain grade of chronic inflammation (e.g. diabetes, obesity) may promote rather unfavorable clinical outcomes. There is an urgent need to further elucidate the molecular and cellular mechanisms involved in progression from mild to life-threatening disease.

Lymphopenia is a near-uniform finding in severe cases of SARS-CoV-2 infection, like in several other viral infections, and typically coincides with profound alterations of the myeloid cell compartment, especially the depletion of CD14 low CD16 high non-classical monocytes and the expansion of the proportion of circulating neutrophils. 30, 31 High expression of T-cell-suppressing molecules programmed death ligand-1 and arginase-1 by neutrophils from COVID-19 patients may further contribute to T-cell malfunction. 30, 32, 33 The elevated NLR that we also observed in our patient population (Table 2) has appeared as an independent biomarker to predict poor clinical outcomes, supporting the notion that neutrophils may play a pivotal role in COVID-19 pathogenesis. 15, 34 Further experimental evidence favoring this hypothesis includes the presence of massive amounts of the neutrophil activation marker S100A8/A9 in the circulation of COVID-19 patients, which correlate positively with neutrophil numbers as well as disease severity and mortality. 31,35-37 Also, the aberrant formation of NETs may contribute to tissue damage and coagulopathy as seen in patients with severe COVID-19. 38, 39 These recent scientific findings sparked our interest to investigate in detail the kinetics of peripheral blood neutrophils from hospitalised COVID-19 patients. For this purpose, we had access to fresh blood samples (samples were processed typically within 30 min of withdrawal) and took advantage of an immunomagnetic purification method that allows for fast isolation of cells to minimise artefacts. The study population included patients who were hospitalised in dedicated general wards or in ICUs ( Table 2) . We compared the parameters with those from age-and sex-matched healthy individuals.

In line with a previously published report describing the presence of immature neutrophils in bronchoalveolar lavage fluid and in the circulation of COVID-19 patients, 31 we detected large numbers of immature neutrophils in the peripheral blood of ICU patients at the time of admission. The occurrence of these immature cells in the periphery indicates emergency myelopoiesis: a haematological response to severe systemic inflammation. 40, 41 The observed high plasma levels of G-CSF may at least partially explain the presence of immature neutrophils in the periphery. Indeed, G-CSF is known to cause a 'shift to the left', that is the occurrence of immature neutrophils in peripheral blood, suggesting a pivotal role for this lineage-specific cytokine in emergency myelopoiesis. 16, 42, 43 Moreover, we detected increased levels of TIMP-1 in plasma from COVID-19 patients. In mice, increased TIMP-1 levels are known to stimulate enrichment of myeloid progenitors and upregulation of genes related to granulopoiesis. 44 Neutrophil activation requires tight regulation to guarantee immune surveillance and to prevent deleterious immune activation leading to tissue damage. 45, 46 Evaluation of the activation state of peripheral blood neutrophils from COVID-19 47, 48 Additional evidence for the presence of highly activated neutrophils in the periphery of patients with severe COVID-19 is provided by the massive protein concentrations of neutrophil elastase, increased gelatinolytic activity in plasma and enhanced concentrations of circulating TIMP-1/MMP-9 complexes. Increased MMP-9 plasma levels were associated with mortality in COVID-19 patients. 37 Moreover, high plasma levels of the archetypal neutrophilattracting chemokine CXCL8 and the myelopoietic cytokine G-CSF may contribute to the activation of peripheral neutrophils. 16, 42, 49 Strikingly, occasional high expression of CXCR3the receptor for the interferon-induced chemokines CXCL9, CXCL10 and CXCL11and CXCR4 was detected on neutrophils from ICU patients (Supplementary figure 2b) . These receptors are not typically found on circulating neutrophils but can be upregulated in an inflamed environment (CXCR3) or appear on immature or aged neutrophils (CXCR4). 48 In addition, enhanced levels of CXCL10 and CXCL11, the two most potent CXCR3 ligands, and reduced activity of their deactivating enzyme CD26 were detected in plasma from COVID-19 patients. Noteworthy, former research efforts confirmed the expression of the CXCR4 gene by neutrophils from COVID-19 patients. 31 In interpreting the overall results of this study, we acknowledge that our sample size for analysis of patient neutrophils is small because of the practical limitations that come with neutrophil experiments, in particular the need of highly fresh blood samples. Moreover, patients were at different treatment strategies and had divergent comorbidities ( Table 2) . Artificial ventilation and other inherent aspects of admittance to ICU may induce changes in neutrophil function and responsiveness that we are unable to anticipate at this moment. Severe COVID-19 has a strong immunopathogenic component, and disease complexity prevents us from understanding to which stimulus the neutrophils are actually responding. In addition, the disease courses of individual patients were highly variable. Surprisingly, no significant differences in neutrophil phenotype or polarisation responses were found between patients with or without treatment with steroids. Moreover, in contrast to most other studies that use whole blood or rely on the accidental presence of neutrophils within the peripheral blood mononuclear cell (PBMC) layer, we analysed highly pure and freshly isolated neutrophils. 31, 50 Collectively, and for the first time, we report the kinetics of peripheral blood neutrophils and their products of activation. We demonstrate increased numbers of immature and activated neutrophils in the circulation from patients with severe COVID-19. These populations disappear towards discharge from ICU. Moreover, we show that SARS-CoV-2 is unable to replicate in human neutrophils in vitro, suggesting that neutrophils are unlikely to support SARS-CoV-2 replication in patients. The lack of evidence indicating infection or direct interaction between SARS-CoV-2 and neutrophils also suggests that phenotypical and functional changes observed in neutrophils from 

Patients were recruited at the University Hospital Leuven. Blood samples were collected from ICU patients during the first 48 h after admission, after one week (between days 6 and 8) and upon discharge from the ICU. In addition, samples were collected from patients upon admission to general wards. Blood samples from age-and sex-matched healthy individuals were investigated for comparative purposes. For analysis of neutrophils and plasma, blood samples were collected in vacutainer tubes treated with ethylenediaminetetraacetic acid (EDTA) or with sodium citrate, respectively (BD Biosciences, Franklin Lakes, NJ) (refer to Table 2 for detailed characteristics of patients). Due to the limited availability of plasma and fresh blood within 30 min of withdrawal, not all patient samples could be included at each time point and in every experiment performed.

Blood samples were spun down for 10 min at 400 g. The supernatant was collected and centrifuged for 20 min at 16 000 g to obtain platelet-free plasma. Plasma was stored until further use at -80°C.

Neutrophils used in shape change assays and for phenotypical characterisation were isolated from the whole blood by immunomagnetic negative selection (EasySep TM Direct Human Neutrophil Isolation Kit; Stemcell Technologies, Vancouver, Canada) within 30 min of withdrawal. Information on neutrophil purity is included in Supplementary figure 3a. Neutrophils used in SARS-CoV-2 exposure assays were isolated from fresh peripheral blood 

Neutrophils were suspended in shape change buffer (1X HBSS without Ca 2+ and Mg 2+ and supplemented with 10 mM HEPES) at a concentration of 0.6 9 10 6 cells mL À1 . The cell suspension (50 µL) was added to 50 µL of chemoattractant dilution and incubated for 0 or 3 min in a flat-bottom 96-well plate followed by fixation with 100 µL Cytofix (BD Biosciences). The following chemoattractants were used: recombinant CXCL8 (72AA; PeproTech, Rocky Hill, NJ, USA; final concentration of 5 or 12.5 ng mL À1 ), recombinant CXCL10 (PeproTech; final concentration of 100 or 300 ng mL À1 ), chemically synthesised CXCL12a (final concentration of 300 ng mL À1 ) 51 and TNF-a (PeproTech; final concentration of 50 ng mL À1 ). Cells were analysed by microscopic evaluation of the cellular shape. Two independent researchers performed the assessment.

Plasma concentrations of G-CSF, GM-CSF, CXCL1, CXCL5, CXCL8, CXCL11 and CXCL12a were measured using customised Meso Scale Discovery (Rockville, MD, USA) multiplex assays. CXCL10 concentrations were evaluated using a specific sandwich ELISA developed in our laboratory. Neutrophil elastase, TIMP-1, TIMP-1/MMP-9 complexes and a2M were quantified by commercially available DuoSet ELISAs (R&D Systems, Minneapolis, MN, USA).

To test metalloproteinase and gelatinase activity, 15 µL of a mixture of dye-quenched (DQ) gelatin (DQ TM Gelatin; Invitrogen, Carlsbad, CA, USA) (final concentration of 3 µg mL À1 ) and OMNIMMP substrate peptide (Mca-PLGL-Dpa-AR-NH 2 , Cat. No. BML-P126-0001; Enzo Life Sciences, Farmingdale, NY, USA) (final concentration of 6 µg mL À1 ) in assay buffer [50 mM Tris, 150 mM NaCl, 5 mM CaCl 2 , 0.01% Tween-20, pH 7.4] was added to 5 µL plasma. Fluorescence was measured over time with the CLARIOstar microplate reader (BMG LABTECH, Ortenberg, Germany). Metalloproteinase activity was inhibited by the addition of 100 mM EDTA. All data shown are representative for the fluorescence measured after 1 h incubation at 37°C. 

The SARS-CoV-2 strain HIAE-02 SARS-CoV-2/SP02/human/ 2020/BRA (GenBank Accession Number MT126808.1), isolated from the second patient diagnosed with COVID-19 in Brazil, was kindly provided by Prof. Edison Luiz Durigon (University of São Paulo, Brazil). SARS-CoV-2 stocks were produced by Vero CCL81 cells. The viral stock titre and identity were assessed by a viral plaque assay and RT-qPCR.

All experiments involving manipulation of infective SARS-CoV-2 were performed in the BSL3 facility of the Laboratory of Emerging Viruses (University of Campinas, Brazil). Purified neutrophils or Vero CCL81 cells were seeded in 24-well plates (1 9 10 6 cells per well) and incubated with SARS-CoV-2 at a multiplicity of infection of 0.1 for 1 h at 37°C and 5% CO 2 . Plates were centrifuged, the viral inoculate was removed and replaced by RPMI medium supplemented with 10% (v/v) FCS and 1% (w/v) penicillin-streptomycin. Samples were collected 6 or 12 h post-infection.

The collected cell supernatants (vide supra) were exposed to UV radiation for 15 min to neutralise viral activity. A mixture containing equal amounts of Color Reagent A and Color Reagent B (R&D Systems) was added to each well of a 96-well plate together with 50 µL of supernatant sample to assess peroxidase activity. Data were acquired at 450 nm in an EnSpire Plate Reader (PerkinElmer, Waltham, MA, USA).

MM obtained a PhD fellowship supported by the L'Or eal -UNESCO for Women in Science Initiative and the FWO-Vlaanderen. SC, MB and LV received PhD fellowships from FWO-Vlaanderen

Mieke Metzemaekers: Conceptualization; Data curation

Formal analysis; Funding acquisition; Investigation

Validation; Visualization; Writing-original draft; Writing-review & editing. Seppe Cambier: Conceptualization; Data curation

Investigation; Methodology; Writing-original draft; Writingreview & editing. Marfa Blanter: Data curation; Formal analysis; Investigation; Methodology; Writing-review & editing. Jennifer Vandooren: Data curation

Investigation; Methodology; Writing-review & editing. Ana Carolina de Carvalho: Formal analysis; Investigation

Writing-review & editing. Bert Malengier-Devlies: Formal analysis; Investigation; Methodology

Writing-review & editing. Lore Vanderbeke: Conceptualization; Data curation; Investigation; Writingreview & editing. Cato Jacobs: Data curation

Writing-review & editing. Sofie Coenen: Data curation

Erik Martens: Formal analysis; Methodology; Writing-review & editing. No€ emie P€ ortner: Formal analysis; Investigation; Writing-review & editing. Lotte Vanbrabant: Formal analysis

Writing-review & editing. Pierre Van Mol: Data curation

Yannick Van Herck: Conceptualization; Data curation; Writing-review & editing. Nathalie Van Aerde: Data curation; Writing-review & editing. Greet Hermans: Data curation; Supervision; Writing-review & editing

Alexandre Borin: Formal analysis; Writing-review & editing. Bruna Toledo N. Pereira: Formal analysis; Writing-review & editing. Arilson Bernardo dos SP Gomes: Formal analysis; Writing-review & editing. St efanie Primon Muraro: Formal analysis

Writing-review & editing. Alessandro S Farias: Data curation

Jos e Luiz Proenca-Modena: Formal analysis; Writing-review & editing. Marco Vinolo: Formal analysis; Writing-review & editing. Contagious Consortium: Data curation; Writing-review & editing. Pedro Elias Marques: Conceptualization; Data curation; Writing-review & editing. Carine Wouters: Conceptualization; Data curation

Supervision; Writing-review & editing. Els Wauters: Conceptualization; Data curation

Investigation; Supervision; Writing-review & editing. Sofie Struyf: Data curation; Formal analysis

Investigation; Supervision; Writing-review & editing. Patrick Matthys: Conceptualization; Data curation

Funding acquisition; Supervision; Writing-review & editing. Ghislain Opdenakker: Conceptualization

Formal analysis; Supervision; Writing-review & editing. Rafael Elias Marques: Conceptualization; Data curation; Formal analysis; Funding acquisition; Investigation; Methodology

Writing-review & editing

Data curation; Funding acquisition; Supervision; Writingreview & editing. Mieke Gouwy: Conceptualization; Data curation; Formal analysis; Investigation; Methodology

Supervision; Writing-review & editing. Paul Proost: Conceptualization; Data curation

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ª 2021 The Authors. Clinical & Translational Immunology published by John Wiley & Sons Australia, Ltd on behalf of Australian and New Zealand Society for Immunology

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Detection of 2019 novel coronavirus (2019-nCoV) by real-time RT-PCR

The assessment of extracellular LDH activity was performed using the LDH Liquiform Kit (Labtest, Lagoa Santa, Brazil) according to the manufacturer's protocol and read at 340 nm using the EnSpire Plate Reader.

RNA was extracted from cell extracts using the Quick-RNA Viral Kit (Zymo Research, Irvine, CA), according to the manufacturer's recommendations. RNA quality and quantity were verified with a NanoDrop One Spectrophotometer (Thermo Fisher Scientific). SARS-CoV-2 RNA quantification was performed by RT-qPCR according to the Charit e protocol 52 using primers and probes for the E gene (forward: 5 0 ACA GGT ACG TTA ATA GTT AAT AGC GT-3 0 , reverse: 5 0 -ATA TTG CAG CAG TAC GCA TAC GCA CAC A-3 0 , probe: 5 0 -6FAM-ACA CTA GCC ATC CTT ACT GCG CTT CG-QSY-3 0 ). All reactions were assembled in a final volume of 12 lL containing 3 lL of TaqMan Fast Virus 1-Step Master Mix (Applied Biosystems, Foster City, CA, USA), 800 nM primers, 400 nM probe and 6 lL of 100-fold diluted RNA in ultrapure water. The cycling algorithm used in this study was as follows: 1 cycle at 50°C for 10 min, 1 cycle at 95°C for 2 min, followed by 45 cycles at 95°C for 5 s and 60°C for 30 s in the QuantStudio3 System (Applied Biosystems). All applicable measures were taken to prevent cross-contamination of samples, and negative and positive controls were included. Reference values for equivalents of plaque-forming unit (ePFU) calculation are Ct means of samples with known viral load (in PFU mL -1 ) plotted as a standard curve. 

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