Background: Neutrophils are the first line of defense against pathogens. They are divided into multiple subpopulations during development and kill pathogens through various mechanisms. Neutrophils are considered one of the markers of severe COVID-19. Summary: In-depth research has revealed that neutrophil subpopulations have multiple complex functions. Different subsets of neutrophils play an important role in the progression of COVID-19. Key Messages: In this review, we provide a detailed overview of the developmental processes of neutrophils at different stages and their recruitment and activation after SARS-CoV-2 infection, aiming to elucidate the changes in neutrophil subpopulations, characteristics, and functions after infection and provide a reference for mechanistic research on neutrophil subpopulations in the context of SARS-CoV-2 infection. In addition, we have also summarized research progress on potential targeted drugs for neutrophil immunotherapy, hoping to provide information that aids the development of therapeutic drugs for the clinical treatment of critically ill COVID-19 patients.

As the first line of defense against pathogens, polymorphonuclear leukocytes (PMNs) respond immediately after pathogen invasion and migrate to the inflamed tissue [1]. PMNs are mainly composed of neutrophils (NEUs) and include a small number of eosinophils and basophils. NEUs are the most abundant leukocyte type in the human body, accounting for 50–70% of all circulating white blood cells, while in mice, they account for only 10–25% [3]. They are produced and released from the bone marrow (BM), and senescent cells are eventually cleared by macrophages in the liver and spleen [4]. The half-life of NEUs is usually only 6–8 h, but they can survive longer under inflammatory conditions [5]. After the last division in the BM, NEUs are released into the blood vessels to migrate freely. Tissue infiltration of NEUs plays a key role in pathogen clearance. Phagocytosis, reactive oxygen species (ROS) generation, and degranulation are classic cell processes used to kill pathogens [6]. Excessive NEU loss can lead to severe immunodeficiency diseases, while excessive activation may cause certain damage to tissues. A current popular research topic is the diverse phenotypes of NEUs, which may play different important functions [7‒9]. Transcriptome and epigenome data have also confirmed that NEUs undergo a series of maturation processes, but their actual function is still a mystery [7].

Coronavirus disease 2019 (COVID-19) is a respiratory disease caused by severe acute respiratory syndrome coronavirus 2 (SARS-CoV-2), with typical clinical symptoms similar to those of influenza, including fever, cough, myalgia, and shortness of breath similar to influenza [11]. Severe cases can develop into severe interstitial pneumonia and even fatal acute respiratory distress syndrome (ARDS) [13]. After infection with the virus, the host’s immune system recognizes SARS-CoV-2 and induces an immune response. Moreover, there is also a substantial change in the proportions of immune cell types. Many studies have shown a higher neutrophil (NEU)-to-lymphocyte ratio (NLR) in COVID-19 patients than in healthy individuals [15]. The NLR is generally considered a marker indicating the severity of COVID-19 and can be used to predict patient prognosis [16‒20]. Various omics analyses have also shown that SARS-CoV-2 infection leads to the appearance of neutrophils in different stages of maturation in the patient’s blood [21‒23]. Neutrophil infiltration in COVID-19 patients was determined through the pathological examination of patient lung tissue [24]. In addition, abundant neutrophil extracellular traps (NETs) were found in the lungs during deceased patient autopsies [25]. Suspected thrombosis related to NETs may be the cause of death [25].

Although the immune response to SARS-CoV-2 has been extensively studied, the role neutrophils play is not clear. In this review, while describing the development, function, and regulation of neutrophil subpopulations, we also emphasize the types of neutrophils found in COVID-19 patients and the impact of neutrophils on the course of disease. In addition, we also summarize some potential neutrophil-targeting agents for the treatment of COVID-19. This review aims to provide new insights into the mechanism of SARS-CoV-2 disease and drug screening.

Developmental Process of Neutrophils

Neutrophils are produced by hematopoietic cord and BM progenitor cells. The traditional developmental stages of neutrophils mainly include myeloblasts, promyelocytes, myelocytes, metamyelocytes, band neutrophils, and mature segmented neutrophils (Fig. 1) [27]. In the metamyelocyte stage, cell division ceases [28]. Neutrophil granule protein gene expression is also associated with this process; Elane and Mpo are expressed in primary granules; Camp, Lcn2, and Ltf are expressed in secondary granules; and Mmp8 and Mmp9 are expressed in tertiary granules [29]. As neutrophils differentiate and mature, they pass through the stem cell pool, mitotic pool, and postmitotic pool in the BM [29].

Fig. 1.

Life cycle of neutrophils in healthy and COVID-19 people. Neutrophils traditionally develop in the BM through myeloblasts, promyelocytes, myelocytes, metamyelocytes, band neutrophils, and mature segmented neutrophils. After SARS-CoV-2 infection, neutrophils are mobilized to lung tissue. There were differences in the neutrophil phenotypes that emerged from mild and severe COVID-19. Neutrophils with immature properties have emerged in severe COVID-19.

Fig. 1.

Life cycle of neutrophils in healthy and COVID-19 people. Neutrophils traditionally develop in the BM through myeloblasts, promyelocytes, myelocytes, metamyelocytes, band neutrophils, and mature segmented neutrophils. After SARS-CoV-2 infection, neutrophils are mobilized to lung tissue. There were differences in the neutrophil phenotypes that emerged from mild and severe COVID-19. Neutrophils with immature properties have emerged in severe COVID-19.

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The production of neutrophils requires the participation of cytokines, such as IL-6, IL-4, and granulocyte colony-stimulating factor (G-CSF). Studies have shown that the number of neutrophils is significantly increased under the stimulation of G-CSF and the regulation of IL-17 and IL-23 [32]. G-CSF can also promote neutrophil mobilization. The different characteristics of G-CSF- and G-CSF receptor-deficient mice have established that G-CSF can regulate granulopoiesis and is a major cytokine affecting neutrophil maturation and mobilization from BM [34]. In addition to G-CSF, CXCR2 signaling also plays an important role in neutrophil mobilization. CXCR2 ligands are mainly responsible for the departure of mature neutrophils from the BM during early acute inflammation, while immature neutrophils typically do not express CXCR2 and do not appear in circulation [36]. CXCR2 ligands induce mobilization much faster than G-CSF, and the mechanisms by which the two induce mobilization may be different [37‒39]. Although both G-CSF and CXCR2 are known to promote neutrophil mobilization, studies have shown that G-CSF may negatively regulate CXCR2-triggered neutrophil mobilization to maintain neutrophil homeostasis and function [36]. Rapid neutrophil mobilization in the early stage of acute inflammation is often facilitated by CXCR2 ligands, but after the acute rapid phase, neutrophil mobilization slows down because G-CSF negatively regulates CXCR2-mediated cell signaling and inhibits neutrophil activation and function [36]. However, not all neutrophils are transported to the peripheral blood and tissues, and most neutrophils remain in the BM. In contrast to CXCR2, CXCL12, a ligand for the chemokine receptor CXCR4 on the neutrophil cell membrane, promotes neutrophil retention in the BM (Fig. 1) [40]. It antagonizes CXCR2 signaling and the two signaling pathways work together to regulate the transport of BM neutrophils [42]. In general, as neutrophils mature, the expression of CXCR2 on their cell membrane surface is upregulated, while the level of CXCR4 is decreased [31].

The process of neutrophil aging is mainly characterized by a decrease in L-selectin (CD62L) levels and the upregulation of CXCR4 and other markers, such as CD11b or CD49d [43]. These aging neutrophils return to the BM after completing their life cycle in the blood and are eliminated in the BM (Fig. 1) [44]. This process may proceed through a CXCR4-dependent mechanism [44‒46]. In the noninflammatory state, circulating neutrophils are preferentially eliminated in the spleen, liver, and BM [47]. In a state of inflammation, neutrophils are eventually recruited to injured tissues and are eliminated by Kupffer cells (liver-resident macrophages) in the vasculature [48]. This process stimulates macrophages to switch to the IL-10hi IL-12lo M2 phenotype [49]. Moreover, the production of IL-23 is reduced, which leads to a decrease in G-CSF release and subsequent neutrophil production [32].

The life cycle of neutrophils has been extensively studied, but the precise delineation of the neutrophil developmental stages remains a challenge. In addition, more precise data are needed on the specific process of CXCR2 signaling-induced neutrophil mobilization.

Neutrophil Subsets

An increasing number of studies have proven that there may be different subsets of neutrophils. The heterogeneity of neutrophils has always been a controversial topic. First, three distinct neutrophil populations, PMN-N, PMN-I, and PMN-II, were identified in mice with different susceptibilities to methicillin-resistant Staphylococcus aureus (MRSA) infection [50]. These neutrophils are classified mainly in terms of the cytokines and chemokines they produce, the effects they have on macrophage activation, and the toll-like receptors (TLRs) and surface antigens they express [50]. In general, neutrophils isolated from MRSA-resistant mice were defined as proinflammatory (IL-12, CCL3), whereas neutrophils from MRSA-susceptible mice were defined as anti-inflammatory (IL-10, CCL2) [50]. A similar phenomenon was observed in tumor-bearing mice [51]. Later studies also demonstrated the existence of distinct lineages of neutrophils. In a study, proinflammatory neutrophils (CD11b+Gr-1+CXCR4low) and proangiogenic neutrophils (MMP9hiCXCR4hi) were recruited to tissues by CXCL2 and vascular endothelial growth factor A (VEGFA), respectively [52]. Kim et al. [53] defined a population of proliferative late-stage neutrophil precursors in the BM through flow cytometry, which was later proven to be highly heterogeneous and present in the BM of both mice and humans [54]. In another study, Evrard et al. [7] using mass cytometry (CyTOF) and whole-transcriptome sequencing (RNA-seq) progressively provided a framework for the process of neutrophil development. Their study showed that the development of neutrophils from neutrophil precursors to mature neutrophils requires an immature neutrophil stage. During the developmental process, gene expression on the surface of neutrophils changes significantly, and their proliferation ability gradually disappears [7]. Immature neutrophils with proliferative ability were C-KIT+CXCR2-CXCR4+, while mature neutrophils highly expressed CXCR2 [7]. In humans, CD10 is a marker to distinguish the maturation state of neutrophils during inflammation. CD66+CD10 neutrophils tend to represent immature neutrophils that lack the immunosuppressive capacity of CD66+CD10+ mature neutrophils [55]. In recent years, single-cell transcriptome profiling has become a powerful tool for finding new subsets of neutrophils. Another group classified neutrophils into G0–G4 immature neutrophils in BM and G5a–G5c mature neutrophils in PB through single-cell transcriptome profiling [56]. By comparison, they found correlations for neutrophils with the definition in previous studies. Pre-neutrophils (pre-Neu), CXCR2lo immature neutrophils (im-Neu), and CXCR2high mature neutrophils (m-Neu) were associated with G2, G3, and G4 neutrophils. Clusters G0, G1, G2, G3, and G4 were aligned with BM GMP, pro-Neu, pre-Neu, immature Neu (imm-Neu), and mature Neu (mNeu), respectively [30].

In severe infections, sepsis, and autoimmune diseases, a population of low-density neutrophils (LDNs) is present [58‒60]. LDNs are often separated by 1.077 g/mL Ficoll gradient centrifugation for clinical analysis. Different from normal-density neutrophils located on top of red blood cells after centrifugation, LDNs are located between the Ficoll and plasma layer [61]. In systemic lupus erythematosus, there are two subsets of LDNs, mature and immature [62]. Previous research has shown that LDNs are closely associated with organ damage and may play a proinflammatory role [63]. In addition, the function of immature LDNs is different from that of typical neutrophils and includes the formation of NETs and degranulation [63]. Blanco-Camarillo et al. [65] demonstrated that LDNs are also present in healthy individuals, albeit in small numbers, and tend to be neutrophils in a mature state. Interestingly, the phenotypes and functions of LDNs and normal-density neutrophils in healthy individuals are so similar that they are almost indistinguishable [65]. In addition, immunosuppressive LDNs have been identified in the peripheral blood of cancer patients, and they are commonly referred to as polymorphonuclear myeloid-derived suppressor cells (PMN-MDSCs) [66‒68]. Myeloid-derived suppressor cells are a heterogeneous population of BM-derived cells that contain neutrophils and monocytes with immunosuppressive functions activated under pathological conditions [66]. PMN-MDSCs have been shown to inhibit T cells, NK cells, and other immunosuppressive functions through a variety of mechanisms [68]. Lectin-type oxidized LDL receptor 1 (LOX-1) has been suggested as a marker to distinguish PMN-MDSCs from neutrophils in cancer patients [27]. However, PMN-MDSCs exhibit phenotypes similar to those of typical neutrophils in mice [27]. Although PMN-MDSCs in humans express CD11b+CD14-CD15+/CD66b+, similar to traditional neutrophils, it has been proven that CD10, CD13, and CD38 can be markers for identifying different maturation stages [74‒76].

Although there are numerous studies on neutrophil subsets, there is still no recognized marker to distinguish them. The functions of different neutrophil subsets may oppose one another. The heterogeneity of immature neutrophils and how they function needs to be further studied. In the future, if the subsets of neutrophils are clearly described, it will make a great contribution to the treatment of many diseases that involve neutrophil biology.

Characteristics of Neutrophils in COVID-19

After the outbreak of COVID-19, notable changes in immune cell populations were documented in COVID-19 patients. Invasion by SARS-CoV-2 induces type I IFN production, as has been shown in the plasma of patients with mild disease. Type I IFN can inhibit viral replication and promote innate and adaptive immunity. Interestingly, the presence of type I IFN has not been found in patients with moderate or severe COVID-19 [77]. A large number of previous studies have shown that compared with healthy people, the number of neutrophils in peripheral blood of severe COVID-19 patients is significantly increased, and a large number of neutrophils also infiltrate in the lung [21]. High levels of calprotectin S100A8 and S100A9 can be found in the plasma of severe COVID-19 patients but not in mild patients [77]. S100A8 and S100A9 are common alarmin proteins that can be released during the formation of NETs or neutrophil death [79]. Through the detection of NETs markers, it can be found that there is a large number of NETs released from the plasma of severe patients. NETs are also found in abundance in the trachea and lung by autopsy of patients and are thought to be associated with the formation of immunothrombosis [25].

At the same time, abnormal immature neutrophils have been observed in severe COVID-19 patients (Fig. 1). In particular, increased CD16low neutrophil numbers were found in patients with moderate and severe COVID-19. Compared with traditional neutrophils, these cells lack the expression of CXCR2, FCG3RB, and CD101, while the expression of CD177, CD11b, and CD62L is decreased, and they highly express CD66b, LOX-1, CD24, and CXCR4, which has the characteristics of immature neutrophils [21]. Other studies have found that neutrophils expressing CEACAM8, ELANE, and LYZ genes can also reflect newly defined neutrophil progenitors and immature neutrophils [80]. In another study, the spontaneous generation of NETs was observed in a group of LDNs expressing CD16int. This group of LDNs exhibits proinflammatory effects and is correlated with coagulopathy and disease severity [81]. Interestingly, conflicting claims have also emerged in different studies regarding the assessment of neutrophil function. Some data have shown abnormal neutrophils with reduced CD62L expression and increased PD-L1 expression in severe CDOID-10 patients, which is reminiscent of PMN-MDSCs with immunosuppressive functions [73] in tumors. However, other studies have shown that PD-L1 expression was not detected in COVID-19 neutrophils [21].

Neutrophils in COVID-19 have also been studied in mouse models. Ly6G is a typical neutrophil marker in mice. In the hACE2 transgenic mouse model, the levels of neutrophils in blood and lung continued to increase after infection [83]. Elevated expression of 100A8 and S100A9 in severe COVID-19 patients was also observed in mice. In the study, mice immunized with SARS-CoV-2, mouse hepatitis virus, influenza A, and other viruses were compared with controls. CD45+CD11b+Ly6Gvariable neutrophils were found in the blood and lungs of SARS-CoV-2-infected mice, whereas only CD45+CD11b+Ly6Ghigh neutrophil populations were present in several other stimulated mice. The authors suggest that the presence of this abnormal cell population represents immature neutrophils [83].

SARS-CoV-2 has evolved many variants from its ancestral strain, and patients infected with these variants also have different neutrophil immune responses. The Omicron strain is currently the most prevalent. In patients with Omicron infection, the neutrophil count and NLR in severe patients were significantly higher than those in mild patients. No significant reductions in these measures were observed in patients with severe Omicron compared with severe COVID-19 patients infected with ancestral strains [84]. Analysis of samples from Omicron convalescent patients by scRNA-seq revealed LDNs with different IL-1β or IFN reactivity. Among them, PI3+ LDNs are considered to have a higher ability to protect against excessive immune damage, and they occur more frequently in recovering patients with mild symptoms. PI3 is an elastase inhibitor that blocks tissue damage caused by excessive immunization [85]. However, CEACAM8+ LDNs with immature properties are more easily activated and exert their effects directly [86]. Compared with the Omicron strain, the other variants have been less studied on neutrophils. There are only a few reports in the literature that in human samples infected with alpha variant, researchers detected significantly higher immature neutrophil count values in severe patients than in mild and moderate patients by automated hematology analyzers [87]. In the mouse model, delayed cytokine release was observed in hACE2 mice infected with Beta strain compared with the ancestral strain. In the late stage of infection, the mice developed severe disease or even death. In severe mice, researchers found significantly increased CXCL1/2 levels, and the mice had even more recruited neutrophils in their lungs than ancestral strain-infected mice [88]. The NLR of patients infected with Delta variant is significantly increased in severe COVID-19, but it did not differ much from the NLR of severe Omicron patients [89]. At present, there is paper that summarizes the effects of two consecutive amino acid mutations R203K and G204R (KR) in nucleocapsid (N) protein on patients and cellular immune responses [91]. The results showed that the neutrophil level and NLR of KR patients were higher than those of ancestral strains patients, and the expression of proinflammatory cytokines, chemokines, and interferon stimulation genes was significantly higher than that of RG patients, which was more likely to cause immune overreaction and lead to severe disease [91].

There are still many mysteries about the appearance of neutrophils in SARS-CoV-2. Some researchers have proposed that the development of neutrophils is altered in the presence of SARS-CoV-2, which results in the appearance of abnormal neutrophils [21]. Some scientists believe that the burst of calprotectin leads to the urgent production of neutrophils in the BM, which are then released into the blood and recruited to the lung tissue of critically ill patients [77]. However, the reason for the appearance of immature neutrophils needs to be explored more. Mouse model is also an important tool to study these neutrophils, but in fact, more marker genes representing different maturation stages of neutrophils in mice may be needed for further study.

Neutrophils are considered immune cells with strong antimicrobial capacity; they can kill microorganisms through phagocytosis, ROS generation, and degranulation that are classic cell processes used to kill pathogens. But the role they play in viral infections has received little research attention [92]. Neutrophils have been shown to be present in respiratory diseases and may cause lung injury [93]. Severe COVID-19 caused by persistent SARS-CoV-2 infection can lead to viral pneumonia and ARDS [95]. In addition, extravasation of neutrophils in the pulmonary capillaries, myocardium, and liver has been observed in COVID-19 patient samples, and microvascular thrombosis was also observed in these sites [96]. As the disease progresses to the severe stage, it has been observed that platelets and neutrophils in COVID-19 patients are significantly overactivated and that the NLR increases [96]. Neutrophilia has been described as an indicator of severe disease and poor prognosis in patients with COVID-19 [20]. Studying the role of neutrophils in COVID-19 progression may provide insights for the prevention and treatment of severe disease.

Neutrophil Recruitment and Migration

The recruitment and migration of neutrophils usually require a multistep pathway that includes binding, rolling, adhesion, crawling, and finally migration. Tethering and rolling are the first steps in the leukocyte recruitment cascade (Fig. 2) [1]. After endothelial cell activation, P-selectin is rapidly upregulated in Weibel-Palade bodies, while E-selectin is synthesized de novo and upregulated within approximately 90 min [98]. After the Weibel-Palade bodies are translocated to the apical cell membrane, both P-selectin and E-selectin bind to their glycosylation ligands, including P-selectin glycoprotein ligand-1 (PSGL-1), E-selectin ligand-1 (ESL-1), CD44, and other glycosylation ligands, to form transmembrane adhesion molecules [100]. This transient adhesion causes the flowing neutrophils to slow down and roll, facilitating their sampling of the local microenvironment to discover pathogen molecules [33]. P-selectin normally mediates fast rolling, whereas E-selectin mediates slow rolling [101]. In addition, L-selectin is also involved in neutrophil binding and rolling, and its main role is to promote neutrophil secondary binding and facilitate neutrophil adhesion or migration [102]. Chemokines anchored by heparan sulfate on the surface of endothelial cells promote neutrophil adhesion and arrest through G-protein-coupled receptor-mediated signaling [104]. During the attachment and rolling of neutrophils on the endothelium, cell-surface integrins interact with ligands (ICAM-1 and ICAM-2) on the inflamed endothelium, leading to conformational changes in integrins. The process of integrin activation is mediated by intracellular signaling molecules such as kindlin-3, talin-1, guanine exchange factors for small GTPases, Rap1 and Rap1-GTP interacting adaptor molecules. This activation eventually leads to a slowing of β2-mediated neutrophil rolling and promotes its arrest along the endothelium [33]. Indeed, neutrophil recruitment is not only activated in a selectin- or integrin-dependent manner [106]. In recent years, dipeptidase 1 (DPEP1), an adhesion receptor found in the liver and lung endothelium, has also been shown to recruit neutrophils, which means that the adhesion cascade can also be carried out in an integrin- and selectin-independent manner [108]. After attachment, neutrophils need to find a site to migrate to and cross the endothelial cell barrier through paracellular (between endothelial cells) or transcellular (through endothelial cells) pathways, which is called diapedesis [109]. After completing the above steps, leukocytes need to pass through the pericyte layer within the venous basement membrane to reach the tissue [111].

Fig. 2.

Neutrophil recruitment cascade in COVID-19. After SARS-CoV-2 invasion, neutrophils are recruited to lung tissue. Under the action of selectin, neutrophils first undergo tethering and rolling. Neutrophil adhesion, crawling, and transmigration mainly require the involvement of integrins. Eventually, neutrophils cross the endothelial cell barrier by paracellular (between endothelial cells) or transcellular (through endothelial cells) pathways. Excessive activation of neutrophils can lead to the production of a large number of cytokines, which may eventually cause lung injury and the formation of immune thrombosis.

Fig. 2.

Neutrophil recruitment cascade in COVID-19. After SARS-CoV-2 invasion, neutrophils are recruited to lung tissue. Under the action of selectin, neutrophils first undergo tethering and rolling. Neutrophil adhesion, crawling, and transmigration mainly require the involvement of integrins. Eventually, neutrophils cross the endothelial cell barrier by paracellular (between endothelial cells) or transcellular (through endothelial cells) pathways. Excessive activation of neutrophils can lead to the production of a large number of cytokines, which may eventually cause lung injury and the formation of immune thrombosis.

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The angiotensin-converting enzyme 2 (ACE2) receptor is a recognized receptor for SARS-CoV-2, and CD147 is involved in SARS-CoV-2 tropism and may be a substitute receptor for ACE2, although there is still a lack of strong evidence [112‒115]. CD147 is expressed in the neutrophils of healthy people, and some studies have shown that it is upregulated in patients with COVID-19, so it may be directly related to SARS CoV-2 infection [21]. Typically, after the fusion of SARS-CoV-2 with the cell membrane, viral RNA is released and replicates (Fig. 2) [116]. Subsequently, immune cells can recognize the pathogen-associated molecular pattern of the virus through pattern recognition receptors (PRRs) [117]. TLRs are one type of PRR and a group of transmembrane proteins [119]. After a TLR recognizes SARS-CoV-2, it can trigger downstream inflammatory responses, such as binding to the adaptor molecule MyD88. A study has shown that neutrophil activity in COVID-19 may be mainly mediated by TLR8 signaling [120]. However, other studies have shown that the spike protein of SARS-CoV-2 can interact with TLR4 to activate immune responses, and TLR4 signaling has been implicated in calprotectin production during SARS-CoV-2 infection [121]. However, the role of TLR4 inhibitors remains to be confirmed in animal and clinical studies. Then, MyD88 activates the transcription factor NF-kB and mitogen-activated protein kinase pathways to induce the release of inflammatory factors, such as chemokines and TNF, IFN, IL-1, and other proinflammatory cytokines, which subsequently induce inflammation and leukocyte accumulation, especially neutrophils [123]. In a study, neutrophils were found to infiltrate the lungs in mice in the early stages of infection [125], and CXCL5 majorly contributes to the recruitment of neutrophils into the lungs during the early stages of infection [125]. In fact, a significant increase in serum proinflammatory cytokine levels and a large amount of neutrophil infiltration were observed in patients with severe COVID-19 [13]. Based on the neutrophil depletion test, it was also found that lung inflammation decreased [125]. In vitro research has shown that NETs generated from neutrophils can promote lung epithelial cell death [26]. Therefore, the excessive recruitment of neutrophils during the immune response of patients with severe COVID-19 may be one of the reasons for severe lung injury (Fig. 2). In addition, the massive release of cytokines may lead to platelet activation, endothelial cell dysfunction, and complement activation, ultimately leading to the formation of fatal immunothrombosis (Fig. 2) [96].

Neutrophil Activation and Degranulation

The functions of neutrophils mainly include phagocytosis, ROS production, degranulation, and NETosis (Fig. 3a). Neutrophils have been confirmed to play an antiviral role in the early stage of viral infection, and activated neutrophils were found to be involved in the IFN-mediated antiviral response in moderate COVID-19 patients [126]. However, degranulation and excessive activation of neutrophils may cause lung injury, and activated neutrophils in the peripheral blood may co-relate with a high risk of death [128]. The granules of neutrophils mainly include primary (azurophilic) granules, secondary (specific) and tertiary (gelatinase) granules, and secretory vesicles, each of which is produced at different stages of neutrophil differentiation (Fig. 1) [7]. Azurophilic granules are first produced during neutrophil development and are classified as peroxisome-positive granules [131]. Their function is primarily to release proteins and peptides that can kill invading pathogens. Secondary or specific granules contain lactoferrin. Tertiary granules contain gelatinases such as matrix metalloproteinase 9 (MMP9). Secretory vesicles release serum albumin as well as presynthesized cytokines [132]. After activation, neutrophils can regulate granule mobilization through two upstream/downstream pathways [132]. The first is adhesion dependent, while the second requires binding to ligands and activating immune receptors. In the process of adhesion, the participation of β2-integrin is needed, and then the signal is transmitted through the Src kinase, Fgr, and Hck cascade signaling pathway [133]. Afterward, immune receptor-ligand interactions and Ca2+ release from the cell occur [134], and as the concentration of Ca2+ increases, neutrophil granules are released in an order opposite to that produced [135].

Fig. 3.

a Neutrophil function and therapeutic targets in COVID-19. There are four functions of neutrophils: phagocytosis, ROS release, degranulation, and NETosis. b Possible pathways through which SARS-CoV-2 causes NETosis. After SARS-CoV-2 invasion, NETs can be stimulated in direct or indirect ways and there are possible therapeutic targets.

Fig. 3.

a Neutrophil function and therapeutic targets in COVID-19. There are four functions of neutrophils: phagocytosis, ROS release, degranulation, and NETosis. b Possible pathways through which SARS-CoV-2 causes NETosis. After SARS-CoV-2 invasion, NETs can be stimulated in direct or indirect ways and there are possible therapeutic targets.

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Multiple studies have shown that myeloperoxidase (MPO) and calprotectin (S100A8/S100A9) are upregulated in the peripheral blood, respiratory tract, and lungs of patients with severe COVID-19 [21]. MPO is the most abundant protein in neutrophils [137]. Although MPO may play an active role in the initial clearance of SARS-CoV-2 infection, the overexpression of MPO may lead to the excessive generation of hypochlorous acid (HOCl) or other oxidants [138]. These oxidants can participate in the production of ROS, including hydrogen peroxide, hydroxyl radicals, HOCl, and singlet oxygen [137]. At present, some studies have investigated the activation pathway of neutrophils by in vitro experiments. Using cell experiments, the authors demonstrated that SARS-CoV-2 virus particles can directly activate neutrophils by activating actin polymerization and binding to PRR, resulting in degranulation and production of IL-8 and ROS [139]. The main function of ROS is to assist in killing microorganisms after being released into the phagosome [6]. Upon the fusion of intracellular particles with phagosomes, neutrophils can recruit NADPH oxidase into phagosomes, and the activation of NADPH oxidase leads to a sharp increase in ROS-associated oxygen consumption [33]. After NADPH oxidase activation, NADPH is rapidly depleted, resulting in molecular oxygen reduction and ROS bursting, which facilitates the catalysis of MPO and H2O2 to produce HOCl [140]. SARS-CoV-2 ancestral strain and different variants were also compared with the ability to stimulate neutrophils to degranulate and generate NETs in vitro. It was finally found that the Delta variant had a significantly increased ability to activate neutrophils compared to the original strain, while Omicron had a reduced ability to activate neutrophils. However, in vitro experiments cannot fully simulate the complex immune response after SASR-CoV-2 invasion [139]. Altogether, dysregulation of these cytotoxic particles and ROS in patients with COVID-19 can cause substantial tissue damage.

NETs

NETs are a network structure composed of histone proteins, elastase, MPO, and cathepsin G and are located on the decondensed chromatin scaffold. Typically, NETosis is used to describe the process of neutrophil production and release of NETs [141]. NETosis can help neutrophils capture and kill pathogens such as bacteria, fungi, and viruses, but when NETs are dysregulated, it may lead to the development of immune-related diseases.

The process of NETosis results in the release of NETs that may cause cell death. Phorbol myristate acetate can be used to artificially activate intracellular pathways in neutrophils to generate NETs. This process involves twenty-four related proteins, including neutrophil elastase (NE, ELANE), MPO, calprotectin, lactotransferrin, and actin [142]. Researchers used phorbol myristate acetate stimulation to explore the role of NETosis and found that NETs were associated with neutrophil death [143]. The increase in ROS production is an important intracellular process leading to NETosis, and NADPH oxidase plays a key role in the ROS pathway in the NETosis process. When NADPH oxidase is activated, it stimulates the production of ROS and the activation of peptide arginine deaminase-4 (PAD4) through the PKC and Raf-MEK-ERK signaling pathways, followed by histone hypercitrullination and chromatin decondensation [144‒146]. PAD4 is a ribozyme that is a key enzyme in NETs formation. After that, elastase (NE) and MPO are activated and transferred from the azophilic granules to the nucleus [144]. Under the joint action of NE and MPO, chromatin decondensation occurs, the nuclear membrane is destroyed, and chromatin is released into the cytoplasm [144]. Therefore, chromatin undergoes further modification by proteins. Finally, NETs destroy the plasma membrane and are released while simultaneously inducing neutrophil death [147].

An alternative pathway for the formation of NETs can occur independently of NADPH oxidase and without cell death, which is termed nonlytic NETosis. This formation pathway is more likely to be associated with infection [148]. After stimulation by Staphylococcus aureus, TLR2 is activated. After a series of processes, such as NE activation and translocation such as suicide NETosis, protein-modified chromatin is expelled through the nuclear membrane and vesicles [149‒151]. At this time, neutrophils still remain active and continue to participate in uptake and phagocytosis. Similarly, NETs can form after the activation of complement by Candida albicans through the nonlytic pathway. NETs formation via TLR4 activation by Escherichia coli or LPS-activated platelets is also achieved by this alternative pathway [152]. The release process of NETs induced by ionomycin and nicotine is similar to that described above [154]. However, the process of NETs production stimulated by ionomycin and nicotine depends on mitochondrial ROS release; that is, NETs production cannot occur completely independently of oxidants [154].

Studies have shown that compared to patients without ARDS, patients with pneumonia-related ARDS contain higher levels of NETs in plasma and bronchial alveolar fluid [156]. In COVID-19 patients, free DNA, MPO-DNA complex, and citrullinated histone H3 levels are significantly increased, and the latter two are markers of NETs, which means that the concentration of NETs in the blood of COVID-19 patients is increased [158]. In general, the increase in plasma NETs is related to the severity of COVID-19 [26]. After SARS-CoV-2 invades respiratory epithelial cells, the cells are stimulated to secrete cytokines, chemokines, and DAMPs. Neutrophils are recruited to the infected site in response to chemokines. Upon activation, neutrophils produce respiratory bursts, releasing ROS and triggering PAD4, NE, and gasdermin D activation, thereby inducing the generation of NETs. In addition, studies have confirmed that NETs are also closely related to the formation of immune thrombosis in the lungs, myocardium, and liver of COVID-19 patients [25]. Moreover, such vascular occlusion caused by NETs has also been found in the liver and kidney [159]. Although NETs are important for defending against invading pathogens, if overproduced, they can lead to tissue damage and autoimmune inflammation and may even induce immune thrombosis when activated.

SARS-CoV-2 can directly induce the formation of NETs and NETosis (Fig. 3b) [26]. Compared to inactivated viruses, live viruses and viruses in a replication state are more likely to stimulate the production of NETs [26]. After SARS-CoV-2 enters host cells through the ACE2/TMPRSS2 pathway, it can directly mediate the occurrence of NETosis in neutrophils, which requires the participation of PAD-4 [26]. In addition, SARS-CoV-2 can invade cells through the C-type lectin receptor [160], which has been shown to induce the formation of NETs, while MPO can directly induce the activation of C3 and participate in the alternative complement pathway, which positively regulates NETosis [160]. In addition, the number of LDNs increased in patients with severe COVID-19, which are more likely to produce NETs and lead to microvascular thrombosis and organ damage [81]. After SARS-CoV-2 invasion, monocytes or macrophages are also activated, which leads to the release of proinflammatory mediators such as IL-1β and IL-18 [163]. These inflammatory cytokines can induce increased NETs production in tissues and blood [163]. Moreover, NETs can induce macrophages to release IL-1β and cause further lung damage [164]. The two processes promote each other and may accelerate the process of immune dysregulation [164]. SARS-CoV-2 can also directly bind to platelet ACE2, induce platelet activation, promote the formation of leukocyte-platelet aggregates, lead to vascular injury and thrombosis, and eventually increase the production of NETs [96]. In addition, NETs can activate tissue factors to trigger the intrinsic coagulation pathway and can also directly activate the contact-dependent coagulation pathway through electrostatic interactions [166]. The histones released by NETs can also further mediate thrombin production [167]. NE and MPO released by NETs can also further aggravate coagulation by eliminating the natural anticoagulants tissue factor pathway inhibitor and thrombomodulin [168]. The combined action of these factors eventually leads to COVID-19-related microvascular thrombosis in tissues, causing endothelial damage and playing an important role in the progression to severe stages of the disease.

With a deeper understanding of the relationship between SARS-CoV-2 and neutrophils, researchers are developing related drugs. At present, many drugs targeting neutrophils start by inhibiting NETs (Fig. 3b). Currently, some NET-targeted drugs have been launched, and more drugs are under development. Examples include inhibitors of histones, elastase NE, PAD4, and gasdermin D, which are required for NETs formation. NE inhibitors can not only block the toxic effect of NE but also inhibit the formation of NETs. Lonodelestat, alvelestat, CHF6333, and elafin have passed phase 1 clinical trials [169‒171]. PAD4 inhibitors (Cl-amidine, YW-56, and GSK484) can also limit the production of NETs, thereby preventing the release of NETs and development of thrombosis in SARS-CoV-2 infection [172]. Among them, Cl-amidine has been proven to inhibit the formation of NETs in patients with COVID-19 and contribute to preventing thrombosis [26]. Studies have shown that disulfiram, a drug used to treat alcoholism, can inhibit the formation of NETs by inhibiting gasdermin D [169]. Ribonuclease α or recombinant human deoxyribonuclease (DNase-1) administered by inhalation has been proven to be involved in thrombolysis and reduce the possibility of thrombosis [173]. Research on the long-acting nanoparticulate DNase-1 formulation in the blood of mice and COVID-19 patients suggests that it may be a potential drug for preventing sepsis in COVID-19 patients [174]. The excessive inflammation caused by immune dysregulation during SARS-CoV-2 infection may ultimately lead to ARDS, thrombosis, and multiorgan damage in patients with severe COVID-19 [96]. Therefore, drugs that alleviate cytokine storms will be beneficial for the treatment of this condition. Colchicine is an existing drug that can inhibit the recruitment of neutrophils to inflammatory sites [175]. Moreover, cytokines such as IL-8, IL-6, and IL-1β can also induce the formation of NETs, so cytokine inhibitors may decrease the formation of NETs [169]. However, clinical trial data on blocking cytokines such as IL-6 are limited [177]. Calprotectin has also been implicated as one of the promoters of cytokine storm disorder. Some investigators have suggested that inhibition of calprotectin with TLR4 inhibitors may be effective in preventing severe COVID-19 manifestations in patients with SARS-CoV-2 variants [122]. At present, there is no specific drug against SARS-CoV-2. The development of drugs targeting neutrophils and NETs may provide new strategies for the treatment of SARS-CoV-2.

Numerous studies have explained the close relationship between neutrophils and SARS-CoV-2 infection. The emergence of immature neutrophil subsets in severe COVID-19 patients also provides a new research direction for the heterogeneity of neutrophils. However, more precise and extensive data are needed to demonstrate the specific function of these heterogeneous neutrophil subpopulations. Much research has been done on the antimicrobial role of neutrophils, but little is known about their antiviral properties. More studies are needed to investigate the effects of degranulation and the release of NETs from activated neutrophils in the antiviral response to infection. In addition, the regulation of neutrophils associated with severe COVID-19 remains a focus for the development of new therapeutic strategies and drugs. How to prevent injury caused by excessive activation of neutrophils while ensuring their normal recruitment at sites of inflammation remains a challenge. In any case, a deeper understanding of neutrophil subsets and functions will greatly help the treatment of severe COVID-19 associated with neutrophil dysregulation.

The authors declare that there are no conflicts of interest.

This work was supported by the National Natural Science Foundation of China (Grant No. 32070543), National Key Research and Development Project of China (Grant No. 2022YFC2303404), and Beijing Natural Science Foundation (Grant No. Z210014).

Na Rong drafted and made the figures. Xiaohui Wei and Jiangning Liu reviewed the manuscript.

1.
Ley
K
,
Laudanna
C
,
Cybulsky
MI
,
Nourshargh
S
.
Getting to the site of inflammation: the leukocyte adhesion cascade updated
.
Nat Rev Immunol
.
2007
;
7
(
9
):
678
89
. .
2.
Sadik
CD
,
Kim
ND
,
Luster
AD
.
Neutrophils cascading their way to inflammation
.
Trends Immunol
.
2011
;
32
(
10
):
452
60
. .
3.
Mestas
J
,
Hughes
CC
.
Of mice and not men: differences between mouse and human immunology
.
J Immunol
.
2004
;
172
(
5
):
2731
8
. .
4.
Furze
RC
,
Rankin
SM
.
The role of the bone marrow in neutrophil clearance under homeostatic conditions in the mouse
.
FASEB J
.
2008
;
22
(
9
):
3111
9
. .
5.
Summers
C
,
Rankin
SM
,
Condliffe
AM
,
Singh
N
,
Peters
AM
,
Chilvers
ER
.
Neutrophil kinetics in health and disease
.
Trends Immunol
.
2010
;
31
(
8
):
318
24
. .
6.
Segal
AW
.
How neutrophils kill microbes
.
Annu Rev Immunol
.
2005
;
23
:
197
223
. .
7.
Evrard
M
,
Kwok
IWH
,
Chong
SZ
,
Teng
KWW
,
Becht
E
,
Chen
J
,
.
Developmental analysis of bone marrow neutrophils reveals populations specialized in expansion, trafficking, and effector functions
.
Immunity
.
2018
;
48
(
2
):
364
79.e8
. .
8.
Deniset
JF
,
Kubes
P
.
Neutrophil heterogeneity: bona fide subsets or polarization states
.
J Leukoc Biol
.
2018
;
103
(
5
):
829
38
. .
9.
Rosales
C
.
Neutrophil: a cell with many roles in inflammation or several cell types
.
Front Physiol
.
2018
;
9
:
113
. .
10.
Grassi
L
,
Pourfarzad
F
,
Ullrich
S
,
Merkel
A
,
Were
F
,
Carrillo-de-Santa-Pau
E
,
.
Dynamics of transcription regulation in human bone marrow myeloid differentiation to mature blood neutrophils
.
Cell Rep
.
2018
;
24
(
10
):
2784
94
. .
11.
Lu
R
,
Zhao
X
,
Li
J
,
Niu
P
,
Yang
B
,
Wu
H
,
.
Genomic characterisation and epidemiology of 2019 novel coronavirus: implications for virus origins and receptor binding
.
Lancet
.
2020
;
395
(
10224
):
565
74
. .
12.
Chen
N
,
Zhou
M
,
Dong
X
,
Qu
J
,
Gong
F
,
Han
Y
,
.
Epidemiological and clinical characteristics of 99 cases of 2019 novel coronavirus pneumonia in Wuhan, China: a descriptive study
.
Lancet
.
2020
;
395
(
10223
):
507
13
. .
13.
Huang
C
,
Wang
Y
,
Li
X
,
Ren
L
,
Zhao
J
,
Hu
Y
,
.
Clinical features of patients infected with 2019 novel coronavirus in Wuhan, China
.
Lancet
.
2020
;
395
(
10223
):
497
506
. .
14.
Jonigk
D
,
Werlein
C
,
Lee
PD
,
Kauczor
HU
,
Länger
F
,
Ackermann
M
.
Pulmonary and systemic pathology in COVID-19—holistic pathological analyses
.
Dtsch Arztebl Int
.
2022
;
119
(
25
):
429
35
. .
15.
Qin
C
,
Zhou
L
,
Hu
Z
,
Zhang
S
,
Yang
S
,
Tao
Y
,
.
Dysregulation of immune response in patients with coronavirus 2019 (COVID-19) in wuhan, China
.
Clin Infect Dis
.
2020
;
71
(
15
):
762
8
. .
16.
Yang
AP
,
Liu
JP
,
Tao
WQ
,
Li
HM
.
The diagnostic and predictive role of NLR, d-NLR and PLR in COVID-19 patients
.
Int Immunopharmacol
.
2020
;
84
:
106504
. .
17.
Karimi
A
,
Shobeiri
P
,
Kulasinghe
A
,
Rezaei
N
.
Novel systemic inflammation markers to predict COVID-19 prognosis
.
Front Immunol
.
2021
;
12
:
741061
. .
18.
Tatum
D
,
Taghavi
S
,
Houghton
A
,
Stover
J
,
Toraih
E
,
Duchesne
J
.
Neutrophil-to-Lymphocyte ratio and outcomes in Louisiana COVID-19 patients
.
Shock
.
2020
;
54
(
5
):
652
8
. .
19.
Seyit
M
,
Avci
E
,
Nar
R
,
Senol
H
,
Yilmaz
A
,
Ozen
M
,
.
Neutrophil to lymphocyte ratio, lymphocyte to monocyte ratio and platelet to lymphocyte ratio to predict the severity of COVID-19
.
Am J Emerg Med
.
2021
;
40
:
110
4
. .
20.
Liu
Y
,
Du
X
,
Chen
J
,
Jin
Y
,
Peng
L
,
Wang
HHX
,
.
Neutrophil-to-lymphocyte ratio as an independent risk factor for mortality in hospitalized patients with COVID-19
.
J Infect
.
2020
;
81
(
1
):
e6
12
. .
21.
Lourda
M
,
Dzidic
M
,
Hertwig
L
,
Bergsten
H
,
Palma Medina
LM
,
Sinha
I
,
.
High-dimensional profiling reveals phenotypic heterogeneity and disease-specific alterations of granulocytes in COVID-19
.
Proc Natl Acad Sci USA
.
2021
;
118
(
40
):
e2109123118
. .
22.
Ren
X
,
Wen
W
,
Fan
X
,
Hou
W
,
Su
B
,
Cai
P
,
.
COVID-19 immune features revealed by a large-scale single-cell transcriptome atlas
.
Cell
.
2021
;
184
(
23
):
5838
913.e19
. .
23.
Rice
CM
,
Lewis
P
,
Ponce-Garcia
FM
,
Gibbs
W
,
Groves
S
,
Cela
D
,
.
Hyperactive immature state and differential CXCR2 expression of neutrophils in severe COVID-19
.
Life Sci Alliance
.
2023
;
6
(
2
):
e202201658
. .
24.
Xu
Z
,
Shi
L
,
Wang
Y
,
Zhang
J
,
Huang
L
,
Zhang
C
,
.
Pathological findings of COVID-19 associated with acute respiratory distress syndrome
.
Lancet Respir Med
.
2020
;
8
(
4
):
420
2
. .
25.
Middleton
EA
,
He
XY
,
Denorme
F
,
Campbell
RA
,
Ng
D
,
Salvatore
SP
,
.
Neutrophil extracellular traps contribute to immunothrombosis in COVID-19 acute respiratory distress syndrome
.
Blood
.
2020
;
136
(
10
):
1169
79
. .
26.
Veras
FP
,
Pontelli
MC
,
Silva
CM
,
Toller-Kawahisa
JE
,
de Lima
M
,
Nascimento
DC
,
.
SARS-CoV-2-triggered neutrophil extracellular traps mediate COVID-19 pathology
.
J Exp Med
.
2020
;
217
(
12
):
e20201129
. .
27.
Veglia
F
,
Perego
M
,
Gabrilovich
D
.
Myeloid-derived suppressor cells coming of age
.
Nat Immunol
.
2018
;
19
(
2
):
108
19
. .
28.
Kondo
M
,
Wagers
AJ
,
Manz
MG
,
Prohaska
SS
,
Scherer
DC
,
Beilhack
GF
,
.
Biology of hematopoietic stem cells and progenitors: implications for clinical application
.
Annu Rev Immunol
.
2003
;
21
:
759
806
. .
29.
Borregaard
N
.
Neutrophils, from marrow to microbes
.
Immunity
.
2010
;
33
(
5
):
657
70
. .
30.
Muench
DE
,
Olsson
A
,
Ferchen
K
,
Pham
G
,
Serafin
RA
,
Chutipongtanate
S
,
.
Mouse models of neutropenia reveal progenitor-stage-specific defects
.
Nature
.
2020
;
582
(
7810
):
109
14
. .
31.
Hidalgo
A
,
Chilvers
ER
,
Summers
C
,
Koenderman
L
.
The neutrophil life cycle
.
Trends Immunol
.
2019
;
40
(
7
):
584
97
. .
32.
Ley
K
,
Smith
E
,
Stark
MA
.
IL-17A-producing neutrophil-regulatory Tn lymphocytes
.
Immunol Res
.
2006
;
34
(
3
):
229
42
. .
33.
Mayadas
TN
,
Cullere
X
,
Lowell
CA
.
The multifaceted functions of neutrophils
.
Annu Rev Pathol
.
2014
;
9
:
181
218
. .
34.
Lévesque
JP
,
Hendy
J
,
Takamatsu
Y
,
Simmons
PJ
,
Bendall
LJ
.
Disruption of the CXCR4/CXCL12 chemotactic interaction during hematopoietic stem cell mobilization induced by GCSF or cyclophosphamide
.
J Clin Invest
.
2003
;
111
(
2
):
187
96
. .
35.
Semerad
CL
,
Christopher
MJ
,
Liu
F
,
Short
B
,
Simmons
PJ
,
Winkler
I
,
.
G-CSF potently inhibits osteoblast activity and CXCL12 mRNA expression in the bone marrow
.
Blood
.
2005
;
106
(
9
):
3020
7
. .
36.
Bajrami
B
,
Zhu
H
,
Kwak
HJ
,
Mondal
S
,
Hou
Q
,
Geng
G
,
.
G-CSF maintains controlled neutrophil mobilization during acute inflammation by negatively regulating CXCR2 signaling
.
J Exp Med
.
2016
;
213
(
10
):
1999
2018
. .
37.
Liu
F
,
Wu
HY
,
Wesselschmidt
R
,
Kornaga
T
,
Link
DC
.
Impaired production and increased apoptosis of neutrophils in granulocyte colony-stimulating factor receptor-deficient mice
.
Immunity
.
1996
;
5
(
5
):
491
501
. .
38.
Lieschke
GJ
,
Grail
D
,
Hodgson
G
,
Metcalf
D
,
Stanley
E
,
Cheers
C
,
.
Mice lacking granulocyte colony-stimulating factor have chronic neutropenia, granulocyte and macrophage progenitor cell deficiency, and impaired neutrophil mobilization
.
Blood
.
1994
;
84
(
6
):
1737
46
. .
39.
Fibbe
WE
,
Pruijt
JF
,
Velders
GA
,
Opdenakker
G
,
van Kooyk
Y
,
Figdor
CG
,
.
Biology of IL-8-induced stem cell mobilization
.
Ann N Y Acad Sci
.
1999
;
872
:
71
82
. .
40.
Nauseef
WM
,
Borregaard
N
.
Neutrophils at work
.
Nat Immunol
.
2014
;
15
(
7
):
602
11
. .
41.
Lawrence
SM
,
Corriden
R
,
Nizet
V
.
The ontogeny of a neutrophil: mechanisms of granulopoiesis and homeostasis
.
Microbiol Mol Biol Rev
.
2018
;
82
(
1
):
e00057-17
. .
42.
Eash
KJ
,
Greenbaum
AM
,
Gopalan
PK
,
Link
DC
.
CXCR2 and CXCR4 antagonistically regulate neutrophil trafficking from murine bone marrow
.
J Clin Invest
.
2010
;
120
(
7
):
2423
31
. .
43.
Hidalgo
A
,
Casanova-Acebes
M
.
Dimensions of neutrophil life and fate
.
Semin Immunol
.
2021
;
57
:
101506
. .
44.
Martin
C
,
Burdon
PC
,
Bridger
G
,
Gutierrez-Ramos
JC
,
Williams
TJ
,
Rankin
SM
.
Chemokines acting via CXCR2 and CXCR4 control the release of neutrophils from the bone marrow and their return following senescence
.
Immunity
.
2003
;
19
(
4
):
583
93
. .
45.
Wang
J
,
Hossain
M
,
Thanabalasuriar
A
,
Gunzer
M
,
Meininger
C
,
Kubes
P
.
Visualizing the function and fate of neutrophils in sterile injury and repair
.
Science
.
2017
;
358
(
6359
):
111
6
. .
46.
Casanova-Acebes
M
,
Pitaval
C
,
Weiss
LA
,
Nombela-Arrieta
C
,
Chèvre
R
,
A-González
N
,
.
Rhythmic modulation of the hematopoietic niche through neutrophil clearance
.
Cell
.
2013
;
153
(
5
):
1025
35
. .
47.
Lum
JJ
,
Bren
G
,
McClure
R
,
Badley
AD
.
Elimination of senescent neutrophils by TNF-related apoptosis-inducing [corrected] ligand
.
J Immunol
.
2005
;
175
(
2
):
1232
8
. .
48.
Shi
J
,
Gilbert
GE
,
Kokubo
Y
,
Ohashi
T
.
Role of the liver in regulating numbers of circulating neutrophils
.
Blood
.
2001
;
98
(
4
):
1226
30
. .
49.
Filardy
AA
,
Pires
DR
,
Nunes
MP
,
Takiya
CM
,
Freire-de-Lima
CG
,
Ribeiro-Gomes
FL
,
.
Proinflammatory clearance of apoptotic neutrophils induces an IL-12(low)IL-10(high) regulatory phenotype in macrophages
.
J Immunol
.
2010
;
185
(
4
):
2044
50
. .
50.
Tsuda
Y
,
Takahashi
H
,
Kobayashi
M
,
Hanafusa
T
,
Herndon
DN
,
Suzuki
F
.
Three different neutrophil subsets exhibited in mice with different susceptibilities to infection by methicillin-resistant Staphylococcus aureus
.
Immunity
.
2004
;
21
(
2
):
215
26
. .
51.
Fridlender
ZG
,
Sun
J
,
Kim
S
,
Kapoor
V
,
Cheng
G
,
Ling
L
,
.
Polarization of tumor-associated neutrophil phenotype by TGF-beta: “N1” versus “N2” TAN
.
Cancer Cell
.
2009
;
16
(
3
):
183
94
. .
52.
Christoffersson
G
,
Vågesjö
E
,
Vandooren
J
,
Lidén
M
,
Massena
S
,
Reinert
RB
,
.
VEGF-A recruits a proangiogenic MMP-9-delivering neutrophil subset that induces angiogenesis in transplanted hypoxic tissue
.
Blood
.
2012
;
120
(
23
):
4653
62
. .
53.
Kim
MH
,
Yang
D
,
Kim
M
,
Kim
SY
,
Kim
D
,
Kang
SJ
.
A late-lineage murine neutrophil precursor population exhibits dynamic changes during demand-adapted granulopoiesis
.
Sci Rep
.
2017
;
7
:
39804
. .
54.
Zhu
YP
,
Padgett
L
,
Dinh
HQ
,
Marcovecchio
P
,
Blatchley
A
,
Wu
R
,
.
Identification of an early unipotent neutrophil progenitor with pro-tumoral activity in mouse and human bone marrow
.
Cell Rep
.
2018
;
24
(
9
):
2329
41.e8
. .
55.
Marini
O
,
Costa
S
,
Bevilacqua
D
,
Calzetti
F
,
Tamassia
N
,
Spina
C
,
.
Mature CD10(+) and immature CD10(-) neutrophils present in G-CSF-treated donors display opposite effects on T cells
.
Blood
.
2017
;
129
(
10
):
1343
56
. .
56.
Xie
X
,
Shi
Q
,
Wu
P
,
Zhang
X
,
Kambara
H
,
Su
J
,
.
Single-cell transcriptome profiling reveals neutrophil heterogeneity in homeostasis and infection
.
Nat Immunol
.
2020
;
21
(
9
):
1119
33
. .
57.
Kwok
I
,
Becht
E
,
Xia
Y
,
Ng
M
,
Teh
YC
,
Tan
L
,
.
Combinatorial single-cell analyses of granulocyte-monocyte progenitor heterogeneity reveals an early uni-potent neutrophil progenitor
.
Immunity
.
2020
;
53
(
2
):
303
18.e5
. .
58.
Silvestre-Roig
C
,
Fridlender
ZG
,
Glogauer
M
,
Scapini
P
.
Neutrophil diversity in health and disease
.
Trends Immunol
.
2019
;
40
(
7
):
565
83
. .
59.
Tay
SH
,
Celhar
T
,
Fairhurst
AM
.
Low-density neutrophils in systemic lupus erythematosus
.
Arthritis Rheumatol
.
2020
;
72
(
10
):
1587
95
. .
60.
Rankin
AN
,
Hendrix
SV
,
Naik
SK
,
Stallings
CL
.
Exploring the role of low-density neutrophils during Mycobacterium tuberculosis infection
.
Front Cell Infect Microbiol
.
2022
;
12
:
901590
. .
61.
Hacbarth
E
,
Kajdacsy-Balla
A
.
Low density neutrophils in patients with systemic lupus erythematosus, rheumatoid arthritis, and acute rheumatic fever
.
Arthritis Rheum
.
1986
;
29
(
11
):
1334
42
. .
62.
Kegerreis
BJ
,
Catalina
MD
,
Geraci
NS
,
Bachali
P
,
Lipsky
PE
,
Grammer
AC
.
Genomic identification of low-density granulocytes and analysis of their role in the pathogenesis of systemic lupus erythematosus
.
J Immunol
.
2019
;
202
(
11
):
3309
17
. .
63.
Mistry
P
,
Nakabo
S
,
O'Neil
L
,
Goel
RR
,
Jiang
K
,
Carmona-Rivera
C
,
.
Transcriptomic, epigenetic, and functional analyses implicate neutrophil diversity in the pathogenesis of systemic lupus erythematosus
.
Proc Natl Acad Sci USA
.
2019
;
116
(
50
):
25222
8
. .
64.
Carlucci
PM
,
Purmalek
MM
,
Dey
AK
,
Temesgen-Oyelakin
Y
,
Sakhardande
S
,
Joshi
AA
,
.
Neutrophil subsets and their gene signature associate with vascular inflammation and coronary atherosclerosis in lupus
.
JCI insight
.
2018
;
3
(
8
):
e99276
. .
65.
Blanco-Camarillo
C
,
Alemán
OR
,
Rosales
C
.
Low-density neutrophils in healthy individuals display a mature primed phenotype
.
Front Immunol
.
2021
;
12
:
672520
. .
66.
Veglia
F
,
Sanseviero
E
,
Gabrilovich
DI
.
Myeloid-derived suppressor cells in the era of increasing myeloid cell diversity
.
Nat Rev Immunol
.
2021
;
21
(
8
):
485
98
. .
67.
Moses
K
,
Brandau
S
.
Human neutrophils: their role in cancer and relation to myeloid-derived suppressor cells
.
Semin Immunol
.
2016
;
28
(
2
):
187
96
. .
68.
Tumino
N
,
Besi
F
,
Martini
S
,
Di Pace
AL
,
Munari
E
,
Quatrini
L
,
.
Polymorphonuclear myeloid-derived suppressor cells are abundant in peripheral blood of cancer patients and suppress natural killer cell anti-tumor activity
.
Front Immunol
.
2021
;
12
:
803014
. .
69.
Poschke
I
,
Kiessling
R
.
On the armament and appearances of human myeloid-derived suppressor cells
.
Clin Immunol
.
2012
;
144
(
3
):
250
68
. .
70.
Nagaraj
S
,
Gabrilovich
DI
.
Regulation of suppressive function of myeloid-derived suppressor cells by CD4+ T cells
.
Semin Cancer Biol
.
2012
;
22
(
4
):
282
8
. .
71.
Yang
Y
,
Li
C
,
Liu
T
,
Dai
X
,
Bazhin
AV
.
Myeloid-derived suppressor cells in tumors: from mechanisms to antigen specificity and microenvironmental regulation
.
Front Immunol
.
2020
;
11
:
1371
. .
72.
Condamine
T
,
Dominguez
GA
,
Youn
JI
,
Kossenkov
AV
,
Mony
S
,
Alicea-Torres
K
,
.
Lectin-type oxidized LDL receptor-1 distinguishes population of human polymorphonuclear myeloid-derived suppressor cells in cancer patients
.
Sci Immunol
.
2016
;
1
(
2
):
aaf8943
. .
73.
Cassetta
L
,
Baekkevold
ES
,
Brandau
S
,
Bujko
A
,
Cassatella
MA
,
Dorhoi
A
,
.
Deciphering myeloid-derived suppressor cells: isolation and markers in humans, mice and non-human primates
.
Cancer Immunol Immunother
.
2019
;
68
(
4
):
687
97
. .
74.
Blazkova
J
,
Gupta
S
,
Liu
Y
,
Gaudilliere
B
,
Ganio
EA
,
Bolen
CR
,
.
Multicenter systems analysis of human blood reveals immature neutrophils in males and during pregnancy
.
J Immunol
.
2017
;
198
(
6
):
2479
88
. .
75.
Karakasheva
TA
,
Waldron
TJ
,
Eruslanov
E
,
Kim
SB
,
Lee
JS
,
O’Brien
S
,
.
CD38-Expressing myeloid-derived suppressor cells promote tumor growth in a murine model of esophageal cancer
.
Cancer Res
.
2015
;
75
(
19
):
4074
85
. .
76.
Bergenfelz
C
,
Leandersson
K
.
The generation and identity of human myeloid-derived suppressor cells
.
Front Oncol
.
2020
;
10
:
109
. .
77.
Silvin
A
,
Chapuis
N
,
Dunsmore
G
,
Goubet
AG
,
Dubuisson
A
,
Derosa
L
,
.
Elevated calprotectin and abnormal myeloid cell subsets discriminate severe from mild COVID-19
.
Cell
.
2020
;
182
(
6
):
1401
18.e18
. .
78.
Mellett
L
,
Khader
SA
.
S100A8/A9 in COVID-19 pathogenesis: impact on clinical outcomes
.
Cytokine Growth Factor Rev
.
2022
;
63
:
90
7
. .
79.
Wang
S
,
Song
R
,
Wang
Z
,
Jing
Z
,
Wang
S
,
Ma
J
.
S100A8/A9 in inflammation
.
Front Immunol
.
2018
;
9
:
1298
. .
80.
Wilk
AJ
,
Rustagi
A
,
Zhao
NQ
,
Roque
J
,
Martínez-Colón
GJ
,
McKechnie
JL
,
.
A single-cell atlas of the peripheral immune response in patients with severe COVID-19
.
Nat Med
.
2020
;
26
(
7
):
1070
6
. .
81.
Morrissey
SM
,
Geller
AE
,
Hu
X
,
Tieri
D
,
Ding
C
,
Klaes
CK
,
.
A specific low-density neutrophil population correlates with hypercoagulation and disease severity in hospitalized COVID-19 patients
.
JCI insight
.
2021
;
6
(
9
):
e148435
. .
82.
Schulte-Schrepping
J
,
Reusch
N
,
Paclik
D
,
Baßler
K
,
Schlickeiser
S
,
Zhang
B
,
.
Severe COVID-19 is marked by a dysregulated myeloid cell compartment
.
Cell
.
2020
;
182
(
6
):
1419
40.e23
. .
83.
Guo
Q
,
Zhao
Y
,
Li
J
,
Liu
J
,
Yang
X
,
Guo
X
,
.
Induction of alarmin S100A8/A9 mediates activation of aberrant neutrophils in the pathogenesis of COVID-19
.
Cell Host Microbe
.
2021
;
29
(
2
):
222
35.e4
. .
84.
Wei
YY
,
Wang
RR
,
Zhang
DW
,
Chen
SH
,
Tan
YY
,
Zhang
WT
,
.
Differential characteristics of patients for hospitalized severe COVID-19 infected by the Omicron variants and wild type of SARS-CoV-2 in China
.
J Inflamm Res
.
2023
;
16
:
3063
78
. .
85.
Hilgendorff
A
,
Parai
K
,
Ertsey
R
,
Juliana Rey-Parra
G
,
Thébaud
B
,
Tamosiuniene
R
,
.
Neonatal mice genetically modified to express the elastase inhibitor elafin are protected against the adverse effects of mechanical ventilation on lung growth
.
Am J Physiol Lung Cell Mol Physiol
.
2012
;
303
(
3
):
L215
27
. .
86.
Li
Z
,
Chen
X
,
Dan
J
,
Hu
T
,
Hu
Y
,
Liu
S
,
.
Innate immune imprints in SARS-CoV-2 Omicron variant infection convalescents
.
Signal Transduct Target Ther
.
2022
;
7
(
1
):
377
. .
87.
Georgakopoulou
VE
,
Makrodimitri
S
,
Triantafyllou
M
,
Samara
S
,
Voutsinas
PM
,
Anastasopoulou
A
,
.
Immature granulocytes: innovative biomarker for SARS-CoV-2 infection
.
Mol Med Rep
.
2022
;
26
(
1
):
217
. .
88.
Lee
S
,
Yoon
GY
,
Lee
SJ
,
Kwon
YC
,
Moon
HW
,
Kim
YJ
,
.
Immunological and pathological peculiarity of severe acute respiratory syndrome coronavirus 2 beta variant
.
Microbiol Spectr
.
2022
;
10
(
5
):
e0237122
. .
89.
Radhakrishnan
N
,
Liu
M
,
Idowu
B
,
Bansari
A
,
Rathi
K
,
Magar
S
,
.
Comparison of the clinical characteristics of SARS-CoV-2 Delta (B.1.617.2) and Omicron (B.1.1.529) infected patients from a single hospitalist service
.
BMC Infect Dis
.
2023
;
23
(
1
):
747
. .
90.
Paranga
TG
,
Pavel-Tanasa
M
,
Constantinescu
D
,
Plesca
CE
,
Petrovici
C
,
Miftode
IL
,
.
Comparison of C-reactive protein with distinct hyperinflammatory biomarkers in association with COVID-19 severity, mortality and SARS-CoV-2 variants
.
Front Immunol
.
2023
;
14
:
1213246
. .
91.
Shuaib
M
,
Adroub
S
,
Mourier
T
,
Mfarrej
S
,
Zhang
H
,
Esau
L
,
.
Impact of the SARS-CoV-2 nucleocapsid 203K/204R mutations on the inflammatory immune response in COVID-19 severity
.
Genome Med
.
2023
;
15
(
1
):
54
. .
92.
Galani
IE
,
Andreakos
E
.
Neutrophils in viral infections: current concepts and caveats
.
J Leukoc Biol
.
2015
;
98
(
4
):
557
64
. .
93.
Abraham
E
.
Neutrophils and acute lung injury
.
Crit Care Med
.
2003
;
31
(
4 Suppl l
):
S195
9
. .
94.
Tse
GM
,
To
KF
,
Chan
PK
,
Lo
AW
,
Ng
KC
,
Wu
A
,
.
Pulmonary pathological features in coronavirus associated severe acute respiratory syndrome (SARS)
.
J Clin Pathol
.
2004
;
57
(
3
):
260
5
. .
95.
Pascarella
G
,
Strumia
A
,
Piliego
C
,
Bruno
F
,
Del Buono
R
,
Costa
F
,
.
COVID-19 diagnosis and management: a comprehensive review
.
J Intern Med
.
2020
;
288
(
2
):
192
206
. .
96.
Nicolai
L
,
Leunig
A
,
Brambs
S
,
Kaiser
R
,
Weinberger
T
,
Weigand
M
,
.
Immunothrombotic dysregulation in COVID-19 pneumonia is associated with respiratory failure and coagulopathy
.
Circulation
.
2020
;
142
(
12
):
1176
89
. .
97.
Cai
J
,
Li
H
,
Zhang
C
,
Chen
Z
,
Liu
H
,
Lei
F
,
.
The neutrophil-to-lymphocyte ratio determines clinical efficacy of corticosteroid therapy in patients with COVID-19
.
Cell Metab
.
2021
;
33
(
2
):
258
69.e3
. .
98.
Petri
B
,
Phillipson
M
,
Kubes
P
.
The physiology of leukocyte recruitment: an in vivo perspective
.
J Immunol
.
2008
;
180
(
10
):
6439
46
. .
99.
Phillipson
M
,
Kubes
P
.
The neutrophil in vascular inflammation
.
Nat Med
.
2011
;
17
(
11
):
1381
90
. .
100.
Impellizzeri
D
,
Cuzzocrea
S
.
Targeting selectins for the treatment of inflammatory diseases
.
Expert Opin Ther Targets
.
2014
;
18
(
1
):
55
67
. .
101.
Kunkel
EJ
,
Ley
K
.
Distinct phenotype of E-selectin-deficient mice. E-selectin is required for slow leukocyte rolling in vivo
.
Circ Res
.
1996
;
79
(
6
):
1196
204
. .
102.
Bargatze
RF
,
Kurk
S
,
Butcher
EC
,
Jutila
MA
.
Neutrophils roll on adherent neutrophils bound to cytokine-induced endothelial cells via L-selectin on the rolling cells
.
J Exp Med
.
1994
;
180
(
5
):
1785
92
. .
103.
Zarbock
A
,
Ley
K
,
McEver
RP
,
Hidalgo
A
.
Leukocyte ligands for endothelial selectins: specialized glycoconjugates that mediate rolling and signaling under flow
.
Blood
.
2011
;
118
(
26
):
6743
51
. .
104.
Massena
S
,
Christoffersson
G
,
Hjertström
E
,
Zcharia
E
,
Vlodavsky
I
,
Ausmees
N
,
.
A chemotactic gradient sequestered on endothelial heparan sulfate induces directional intraluminal crawling of neutrophils
.
Blood
.
2010
;
116
(
11
):
1924
31
. .
105.
Margraf
A
,
Ley
K
,
Zarbock
A
.
Neutrophil recruitment: from model systems to tissue-specific patterns
.
Trends Immunol
.
2019
;
40
(
7
):
613
34
. .
106.
Maas
SL
,
Soehnlein
O
,
Viola
JR
.
Organ-specific mechanisms of transendothelial neutrophil migration in the lung, liver, kidney, and aorta
.
Front Immunol
.
2018
;
9
:
2739
. .
107.
Fan
Z
,
McArdle
S
,
Marki
A
,
Mikulski
Z
,
Gutierrez
E
,
Engelhardt
B
,
.
Neutrophil recruitment limited by high-affinity bent β2 integrin binding ligand in cis
.
Nat Commun
.
2016
;
7
:
12658
. .
108.
Choudhury
SR
,
Babes
L
,
Rahn
JJ
,
Ahn
BY
,
Goring
KAR
,
King
JC
,
.
Dipeptidase-1 is an adhesion receptor for neutrophil recruitment in lungs and liver
.
Cell
.
2019
;
178
(
5
):
1205
21.e17
. .
109.
Muller
WA
.
Transendothelial migration: unifying principles from the endothelial perspective
.
Immunol Rev
.
2016
;
273
(
1
):
61
75
. .
110.
Carman
CV
.
Mechanisms for transcellular diapedesis: probing and pathfinding by “invadosome-like protrusions”
.
J Cell Sci
.
2009
;
122
(
Pt 17
):
3025
35
. .
111.
Nourshargh
S
,
Hordijk
PL
,
Sixt
M
.
Breaching multiple barriers: leukocyte motility through venular walls and the interstitium
.
Nat Rev Mol Cell Biol
.
2010
;
11
(
5
):
366
78
. .
112.
Jackson
CB
,
Farzan
M
,
Chen
B
,
Choe
H
.
Mechanisms of SARS-CoV-2 entry into cells
.
Nat Rev Mol Cell Biol
.
2022
;
23
(
1
):
3
20
. .
113.
Fenizia
C
,
Galbiati
S
,
Vanetti
C
,
Vago
R
,
Clerici
M
,
Tacchetti
C
,
.
SARS-CoV-2 entry: at the crossroads of CD147 and ACE2
.
Cells
.
2021
;
10
(
6
):
1434
. .
114.
Wang
K
,
Chen
W
,
Zhang
Z
,
Deng
Y
,
Lian
JQ
,
Du
P
,
.
CD147-spike protein is a novel route for SARS-CoV-2 infection to host cells
.
Signal Transduct Target Ther
.
2020
;
5
(
1
):
283
. .
115.
Shilts
J
,
Crozier
TWM
,
Greenwood
EJD
,
Lehner
PJ
,
Wright
GJ
.
No evidence for basigin/CD147 as a direct SARS-CoV-2 spike binding receptor
.
Sci Rep
.
2021
;
11
(
1
):
413
. .
116.
Gheblawi
M
,
Wang
K
,
Viveiros
A
,
Nguyen
Q
,
Zhong
JC
,
Turner
AJ
,
.
Angiotensin-converting enzyme 2: SARS-CoV-2 receptor and regulator of the renin-angiotensin system: celebrating the 20th anniversary of the discovery of ACE2
.
Circ Res
.
2020
;
126
(
10
):
1456
74
. .
117.
Hou
YJ
,
Okuda
K
,
Edwards
CE
,
Martinez
DR
,
Asakura
T
,
Dinnon
KH
3rd
,
.
SARS-CoV-2 reverse genetics reveals a variable infection gradient in the respiratory tract
.
Cell
.
2020
;
182
(
2
):
429
46.e14
. .
118.
Sungnak
W
,
Huang
N
,
Bécavin
C
,
Berg
M
,
Queen
R
,
Litvinukova
M
,
.
SARS-CoV-2 entry factors are highly expressed in nasal epithelial cells together with innate immune genes
.
Nat Med
.
2020
;
26
(
5
):
681
7
. .
119.
Akira
S
,
Takeda
K
.
Toll-like receptor signalling
.
Nat Rev Immunol
.
2004
;
4
(
7
):
499
511
. .
120.
Gardiman
E
,
Bianchetto-Aguilera
F
,
Gasperini
S
,
Tiberio
L
,
Scandola
M
,
Lotti
V
,
.
SARS-CoV-2-Associated ssRNAs activate human neutrophils in a TLR8-dependent fashion
.
Cells
.
2022
;
11
(
23
):
3785
. .
121.
Zhao
Y
,
Kuang
M
,
Li
J
,
Zhu
L
,
Jia
Z
,
Guo
X
,
.
SARS-CoV-2 spike protein interacts with and activates TLR41
.
Cell Res
.
2021
;
31
(
7
):
818
20
. .
122.
Loh
JT
,
Teo
JKH
,
Lam
KP
.
Dok3 restrains neutrophil production of calprotectin during TLR4 sensing of SARS-CoV-2 spike protein
.
Front Immunol
.
2022
;
13
:
996637
. .
123.
Cao
X
.
COVID-19: immunopathology and its implications for therapy
.
Nat Rev Immunol
.
2020
;
20
(
5
):
269
70
. .
124.
Akira
S
,
Uematsu
S
,
Takeuchi
O
.
Pathogen recognition and innate immunity
.
Cell
.
2006
;
124
(
4
):
783
801
. .
125.
Liang
Y
,
Li
H
,
Li
J
,
Yang
ZN
,
Li
JL
,
Zheng
HW
,
.
Role of neutrophil chemoattractant CXCL5 in SARS-CoV-2 infection-induced lung inflammatory innate immune response in an in vivo hACE2 transfection mouse model
.
Zool Res
.
2020
;
41
(
6
):
621
31
. .
126.
Camp
JV
,
Jonsson
CB
.
A role for neutrophils in viral respiratory disease
.
Front Immunol
.
2017
;
8
:
550
. .
127.
Rosa
BA
,
Ahmed
M
,
Singh
DK
,
Choreño-Parra
JA
,
Cole
J
,
Jiménez-Álvarez
LA
,
.
IFN signaling and neutrophil degranulation transcriptional signatures are induced during SARS-CoV-2 infection
.
Commun Biol
.
2021
;
4
(
1
):
290
. .
128.
Meizlish
ML
,
Pine
AB
,
Bishai
JD
,
Goshua
G
,
Nadelmann
ER
,
Simonov
M
,
.
A neutrophil activation signature predicts critical illness and mortality in COVID-19
.
Blood Adv
.
2021
;
5
(
5
):
1164
77
. .
129.
Dennison
D
,
Al Khabori
M
,
Al Mamari
S
,
Aurelio
A
,
Al Hinai
H
,
Al Maamari
K
,
.
Circulating activated neutrophils in COVID-19: an independent predictor for mechanical ventilation and death
.
Int J Infect Dis
.
2021
;
106
:
155
9
. .
130.
Cowland
JB
,
Borregaard
N
.
Granulopoiesis and granules of human neutrophils
.
Immunol Rev
.
2016
;
273
(
1
):
11
28
. .
131.
Bainton
DF
,
Ullyot
JL
,
Farquhar
MG
.
The development of neutrophilic polymorphonuclear leukocytes in human bone marrow
.
J Exp Med
.
1971
;
134
(
4
):
907
34
. .
132.
Sheshachalam
A
,
Srivastava
N
,
Mitchell
T
,
Lacy
P
,
Eitzen
G
.
Granule protein processing and regulated secretion in neutrophils
.
Front Immunol
.
2014
;
5
:
448
. .
133.
Lacy
P
.
Mechanisms of degranulation in neutrophils
.
Allergy Asthma Clin Immunol
.
2006
;
2
(
3
):
98
108
. .
134.
Futosi
K
,
Fodor
S
,
Mócsai
A
.
Neutrophil cell surface receptors and their intracellular signal transduction pathways
.
Int Immunopharmacol
.
2013
;
17
(
3
):
638
50
. .
135.
Sengeløv
H
,
Kjeldsen
L
,
Borregaard
N
.
Control of exocytosis in early neutrophil activation
.
J Immunol
.
1993
;
150
(
4
):
1535
43
. .
136.
Chiu
KH
,
Yip
CC
,
Poon
RW
,
Leung
KH
,
Li
X
,
Hung
IF
,
.
Correlations of Myeloperoxidase (MPO), Adenosine deaminase (ADA), C-C motif chemokine 22 (CCL22), Tumour necrosis factor alpha (TNFα) and Interleukin-6 (IL-6) mRNA expression in the nasopharyngeal specimens with the diagnosis and severity of SARS-CoV-2 infections
.
Emerg Microbes Infect
.
2023
;
12
(
1
):
2157338
. .
137.
Klebanoff
SJ
.
Myeloperoxidase: friend and foe
.
J Leukoc Biol
.
2005
;
77
(
5
):
598
625
. .
138.
Akgun
E
,
Tuzuner
MB
,
Sahin
B
,
Kilercik
M
,
Kulah
C
,
Cakiroglu
HN
,
.
Proteins associated with neutrophil degranulation are upregulated in nasopharyngeal swabs from SARS-CoV-2 patients
.
PLoS One
.
2020
;
15
(
10
):
e0240012
. .
139.
Lebourgeois
S
,
David
A
,
Chenane
HR
,
Granger
V
,
Menidjel
R
,
Fidouh
N
,
.
Differential activation of human neutrophils by SARS-CoV-2 variants of concern
.
Front Immunol
.
2022
;
13
:
1010140
. .
140.
Odobasic
D
,
Kitching
AR
,
Holdsworth
SR
.
Neutrophil-mediated regulation of innate and adaptive immunity: the role of myeloperoxidase
.
J Immunol Res
.
2016
;
2016
:
2349817
. .
141.
Brinkmann
V
,
Reichard
U
,
Goosmann
C
,
Fauler
B
,
Uhlemann
Y
,
Weiss
DS
,
.
Neutrophil extracellular traps kill bacteria
.
Science
.
2004
;
303
(
5663
):
1532
5
. .
142.
Urban
CF
,
Ermert
D
,
Schmid
M
,
Abu-Abed
U
,
Goosmann
C
,
Nacken
W
,
.
Neutrophil extracellular traps contain calprotectin, a cytosolic protein complex involved in host defense against Candida albicans
.
PLoS Pathog
.
2009
;
5
(
10
):
e1000639
. .
143.
Fuchs
TA
,
Abed
U
,
Goosmann
C
,
Hurwitz
R
,
Schulze
I
,
Wahn
V
,
.
Novel cell death program leads to neutrophil extracellular traps
.
J Cel Biol
.
2007
;
176
(
2
):
231
41
. .
144.
Papayannopoulos
V
,
Metzler
KD
,
Hakkim
A
,
Zychlinsky
A
.
Neutrophil elastase and myeloperoxidase regulate the formation of neutrophil extracellular traps
.
J Cel Biol
.
2010
;
191
(
3
):
677
91
. .
145.
Wang
Y
,
Li
M
,
Stadler
S
,
Correll
S
,
Li
P
,
Wang
D
,
.
Histone hypercitrullination mediates chromatin decondensation and neutrophil extracellular trap formation
.
J Cel Biol
.
2009
;
184
(
2
):
205
13
. .
146.
Li
P
,
Li
M
,
Lindberg
MR
,
Kennett
MJ
,
Xiong
N
,
Wang
Y
.
PAD4 is essential for antibacterial innate immunity mediated by neutrophil extracellular traps
.
J Exp Med
.
2010
;
207
(
9
):
1853
62
. .
147.
Metzler
KD
,
Goosmann
C
,
Lubojemska
A
,
Zychlinsky
A
,
Papayannopoulos
V
.
A myeloperoxidase-containing complex regulates neutrophil elastase release and actin dynamics during NETosis
.
Cell Rep
.
2014
;
8
(
3
):
883
96
. .
148.
Slaba
I
,
Wang
J
,
Kolaczkowska
E
,
McDonald
B
,
Lee
WY
,
Kubes
P
.
Imaging the dynamic platelet-neutrophil response in sterile liver injury and repair in mice
.
Hepatology
.
2015
;
62
(
5
):
1593
605
. .
149.
Pilsczek
FH
,
Salina
D
,
Poon
KK
,
Fahey
C
,
Yipp
BG
,
Sibley
CD
,
.
A novel mechanism of rapid nuclear neutrophil extracellular trap formation in response to Staphylococcus aureus
.
J Immunol
.
2010
;
185
(
12
):
7413
25
. .
150.
Yipp
BG
,
Petri
B
,
Salina
D
,
Jenne
CN
,
Scott
BN
,
Zbytnuik
LD
,
.
Infection-induced NETosis is a dynamic process involving neutrophil multitasking in vivo
.
Nat Med
.
2012
;
18
(
9
):
1386
93
. .
151.
Jorch
SK
,
Kubes
P
.
An emerging role for neutrophil extracellular traps in noninfectious disease
.
Nat Med
.
2017
;
23
(
3
):
279
87
. .
152.
Byrd
AS
,
O’Brien
XM
,
Johnson
CM
,
Lavigne
LM
,
Reichner
JS
.
An extracellular matrix-based mechanism of rapid neutrophil extracellular trap formation in response to Candida albicans
.
J Immunol
.
2013
;
190
(
8
):
4136
48
. .
153.
McDonald
B
,
Urrutia
R
,
Yipp
BG
,
Jenne
CN
,
Kubes
P
.
Intravascular neutrophil extracellular traps capture bacteria from the bloodstream during sepsis
.
Cell Host Microbe
.
2012
;
12
(
3
):
324
33
. .
154.
Douda
DN
,
Khan
MA
,
Grasemann
H
,
Palaniyar
N
.
SK3 channel and mitochondrial ROS mediate NADPH oxidase-independent NETosis induced by calcium influx
.
Proc Natl Acad Sci USA
.
2015
;
112
(
9
):
2817
22
. .
155.
Hosseinzadeh
A
,
Thompson
PR
,
Segal
BH
,
Urban
CF
.
Nicotine induces neutrophil extracellular traps
.
J Leukoc Biol
.
2016
;
100
(
5
):
1105
12
. .
156.
Bendib
I
,
de Chaisemartin
L
,
Granger
V
,
Schlemmer
F
,
Maitre
B
,
Hüe
S
,
.
Neutrophil extracellular traps are elevated in patients with pneumonia-related acute respiratory distress syndrome
.
Anesthesiology
.
2019
;
130
(
4
):
581
91
. .
157.
Lv
X
,
Wen
T
,
Song
J
,
Xie
D
,
Wu
L
,
Jiang
X
,
.
Extracellular histones are clinically relevant mediators in the pathogenesis of acute respiratory distress syndrome
.
Respir Res
.
2017
;
18
(
1
):
165
. .
158.
Zuo
Y
,
Yalavarthi
S
,
Shi
H
,
Gockman
K
,
Zuo
M
,
Madison
JA
,
.
Neutrophil extracellular traps in COVID-19
.
JCI Insight
.
2020
;
5
(
11
):
e138999
. .
159.
Leppkes
M
,
Knopf
J
,
Naschberger
E
,
Lindemann
A
,
Singh
J
,
Herrmann
I
,
.
Vascular occlusion by neutrophil extracellular traps in COVID-19
.
EBioMedicine
.
2020
;
58
:
102925
. .
160.
Lempp
FA
,
Soriaga
LB
,
Montiel-Ruiz
M
,
Benigni
F
,
Noack
J
,
Park
YJ
,
.
Lectins enhance SARS-CoV-2 infection and influence neutralizing antibodies
.
Nature
.
2021
;
598
(
7880
):
342
7
. .
161.
de Bont
CM
,
Boelens
WC
,
Pruijn
GJM
.
NETosis, complement, and coagulation: a triangular relationship
.
Cell Mol Immunol
.
2019
;
16
(
1
):
19
27
. .
162.
Cabrera
LE
,
Pekkarinen
PT
,
Alander
M
,
Nowlan
KHA
,
Nguyen
NA
,
Jokiranta
S
,
.
Characterization of low-density granulocytes in COVID-19
.
PLoS Pathog
.
2021
;
17
(
7
):
e1009721
. .
163.
Park
JH
,
Lee
HK
.
Re-analysis of single cell transcriptome reveals that the NR3C1-CXCL8-neutrophil Axis determines the severity of COVID-19
.
Front Immunol
.
2020
;
11
:
2145
. .
164.
Chousterman
BG
,
Swirski
FK
,
Weber
GF
.
Cytokine storm and sepsis disease pathogenesis
.
Semin Immunopathol
.
2017
;
39
(
5
):
517
28
. .
165.
Zhang
S
,
Liu
Y
,
Wang
X
,
Yang
L
,
Li
H
,
Wang
Y
,
.
SARS-CoV-2 binds platelet ACE2 to enhance thrombosis in COVID-19
.
J Hematol Oncol
.
2020
;
13
(
1
):
120
. .
166.
Kambas
K
,
Chrysanthopoulou
A
,
Vassilopoulos
D
,
Apostolidou
E
,
Skendros
P
,
Girod
A
,
.
Tissue factor expression in neutrophil extracellular traps and neutrophil derived microparticles in antineutrophil cytoplasmic antibody associated vasculitis may promote thromboinflammation and the thrombophilic state associated with the disease
.
Ann Rheum Dis
.
2014
;
73
(
10
):
1854
63
. .
167.
Gould
TJ
,
Vu
TT
,
Swystun
LL
,
Dwivedi
DJ
,
Mai
SH
,
Weitz
JI
,
.
Neutrophil extracellular traps promote thrombin generation through platelet-dependent and platelet-independent mechanisms
.
Arterioscler Thromb Vasc Biol
.
2014
;
34
(
9
):
1977
84
. .
168.
Massberg
S
,
Grahl
L
,
von Bruehl
ML
,
Manukyan
D
,
Pfeiler
S
,
Goosmann
C
,
.
Reciprocal coupling of coagulation and innate immunity via neutrophil serine proteases
.
Nat Med
.
2010
;
16
(
8
):
887
96
. .
169.
Barnes
BJ
,
Adrover
JM
,
Baxter-Stoltzfus
A
,
Borczuk
A
,
Cools-Lartigue
J
,
Crawford
JM
,
.
Targeting potential drivers of COVID-19: neutrophil extracellular traps
.
J Exp Med
.
2020
;
217
(
6
):
e20200652
. .
170.
Yaqinuddin
A
,
Kashir
J
.
Novel therapeutic targets for SARS-CoV-2-induced acute lung injury: targeting a potential IL-1β/neutrophil extracellular traps feedback loop
.
Med Hypotheses
.
2020
;
143
:
109906
. .
171.
Chiang
CC
,
Korinek
M
,
Cheng
WJ
,
Hwang
TL
.
Targeting neutrophils to treat acute respiratory distress syndrome in coronavirus disease
.
Front Pharmacol
.
2020
;
11
:
572009
. .
172.
Elliott
W
Jr
,
Guda
MR
,
Asuthkar
S
,
Teluguakula
N
,
Prasad
DVR
,
Tsung
AJ
,
.
PAD inhibitors as a potential treatment for SARS-CoV-2 immunothrombosis
.
Biomedicines
.
2021
;
9
(
12
):
1867
. .
173.
Park
HH
,
Park
W
,
Lee
YY
,
Kim
H
,
Seo
HS
,
Choi
DW
,
.
Bioinspired DNase-I-coated melanin-like nanospheres for modulation of infection-associated NETosis dysregulation
.
Adv Sci
.
2020
;
7
(
23
):
2001940
. .
174.
Lee
YY
,
Park
HH
,
Park
W
,
Kim
H
,
Jang
JG
,
Hong
KS
,
.
Long-acting nanoparticulate DNase-1 for effective suppression of SARS-CoV-2-mediated neutrophil activities and cytokine storm
.
Biomaterials
.
2021
;
267
:
120389
. .
175.
Dhawan
UK
,
Bhattacharya
P
,
Narayanan
S
,
Manickam
V
,
Aggarwal
A
,
Subramanian
M
.
Hypercholesterolemia impairs clearance of neutrophil extracellular traps and promotes inflammation and atherosclerotic plaque progression
.
Arterioscler Thromb Vasc Biol
.
2021
;
41
(
10
):
2598
615
. .
176.
Yang
L
,
Liu
L
,
Zhang
R
,
Hong
J
,
Wang
Y
,
Wang
J
,
.
IL-8 mediates a positive loop connecting increased neutrophil extracellular traps (NETs) and colorectal cancer liver metastasis
.
J Cancer
.
2020
;
11
(
15
):
4384
96
. .
177.
Hermine
O
,
Mariette
X
,
Tharaux
PL
,
Resche-Rigon
M
,
Porcher
R
,
Ravaud
P
,
.
Effect of tocilizumab vs usual care in adults hospitalized with COVID-19 and moderate or severe pneumonia: a randomized clinical trial
.
JAMA Intern Med
.
2021
;
181
(
1
):
32
40
. .
178.
Salvarani
C
,
Dolci
G
,
Massari
M
,
Merlo
DF
,
Cavuto
S
,
Savoldi
L
,
.
Effect of tocilizumab vs standard care on clinical worsening in patients hospitalized with COVID-19 pneumonia: a randomized clinical trial
.
JAMA Intern Med
.
2021
;
181
(
1
):
24
31
. .