key: cord-1005988-3kmb0xcp
authors: Marcos‐Jiménez, Ana; Sánchez‐Alonso, Santiago; Alcaraz‐Serna, Ana; Esparcia, Laura; López‐Sanz, Celia; Sampedro‐Núñez, Miguel; Mateu‐Albero, Tamara; Sánchez‐Cerrillo, Ildefonso; Martínez‐Fleta, Pedro; Gabrie, Ligia; Guerola, Luciana del Campo; Rodríguez‐Frade, José Miguel; Casasnovas, José M.; Reyburn, Hugh T.; Valés‐Gómez, Mar; López‐Trascasa, Margarita; Martín‐Gayo, Enrique; Calzada, María José; Castañeda, Santos; de la Fuente, Hortensia; González‐Álvaro, Isidoro; Sánchez‐Madrid, Francisco; Muñoz‐Calleja, Cecilia; Alfranca, Arantzazu
title: Deregulated cellular circuits driving immunoglobulins and complement consumption associate with the severity of COVID‐19 patients
date: 2020-11-30
journal: Eur J Immunol
DOI: 10.1002/eji.202048858
sha: 0e8a229c545f115a3e3608c99731cbe9cc794caa
doc_id: 1005988
cord_uid: 3kmb0xcp

SARS‐CoV‐2 infection causes an abrupt response by the host immune system, which is largely responsible for the outcome of COVID‐19. We investigated whether the specific immune responses in the peripheral blood of 276 patients associated to severity and progression of COVID‐19. At admission, dramatic lymphopenia of T, B and NK cells associated to severity. Conversely, the proportion of B cells, plasmablasts, circulating follicular helper T cells (cTfh) and CD56‐CD16+ NK‐cells increased. Regarding humoral immunity, levels of IgM, IgA and IgG were unaffected, but when degrees of severity were considered, IgG was lower in severe patients. Compared to healthy donors, complement C3 and C4 protein levels were higher in mild and moderate, but not in severe patients, while the activation peptide of C5 (C5a) increased from the admission in every patient, regardless their severity. Moreover, total IgG, the IgG1 and IgG3 isotypes and C4 decreased from day 0 to day 10 in patients who were hospitalized for more than two weeks, but not in patients who were discharged earlier. Our study provides important clues to understand the immune response observed in COVID‐19 patients, associating severity with an imbalanced humoral response and identifying new targets for therapeutic intervention. This article is protected by copyright. All rights reserved

Novel coronavirus disease (COVID-19), due to severe acute respiratory coronavirus 2 (SARS-CoV-2), is either asymptomatic or presents with mild symptoms in a majority of individuals. However, up to 20% patients develop a severe form of the disease with pneumonia, which in some cases results in acute respiratory distress syndrome (ARDS) and requires invasive mechanical ventilation. ARDS, together with myocardial damage, are main causes of mortality in COVID-19 [1] .

A pathogenic hallmark of ARDS is the disruption of the alveolar-capillary barrier and a subsequent increase in permeability, which has been partially attributed to a maladaptive immune response.

Thus, lung alveolar macrophages may be infected by the virus and become activated, leading to a cytokine release syndrome, which contributes to endothelial injury with the recruitment and activation of innate and adaptive immune cells and the extravasation of plasma components of the humoral immunity [2] .

Humoral immunity plays a key role in the initial control of viral infections and cell-to-cell spread. It is mediated by the complement system and the immunoglobulins (Ig) [3, 4] . Different SARS-CoV-2 components such as pathogen-associated molecular patterns (PAMPs) and N protein, together with C reactive protein (CRP) from plasma, may activate either the alternative or the lectin pathways of the complement cascade early during infection [5] . Furthermore, early IgM isotype antibodies efficiently trigger the classical complement cascade. Secreted IgA antibodies neutralize viruses within the mucosa of the respiratory and gastrointestinal tracts. Finally, in a more advanced stage of the disease, virus-specific IgG antibodies opsonize viral particles. This may lead to the formation of immune complexes that contribute to the activation of the classical complement pathway.

Furthermore, opsonized viral particles can attach to Fc receptors on phagocytes and NK cells. The latter constitute major innate immunity mediators during anti-viral responses, since they kill infected cells through different mechanisms, including antibody-dependent cell cytotoxicity (ADCC), which is mediated by the FcRIIIA (CD16) binding to clustered IgG displayed on cell surface of virally infected cells [6] . Isotype switching and affinity antibody maturation, as well as the generation of memory B cells and long-lived plasma cells, are B cell responses driven by helper T cells, in particular follicular helper T cells (Tfh).

However, deregulated humoral immune response can damage host tissues. Complement-mediated tissue injury is elicited by an intense inflammatory loop secondary to C3a-and C5a-mediated recruitment and activation of neutrophils, monocytes, macrophages, lymphocytes, and platelets.

Phagocytes in turn generate reactive oxygen species (ROS) and proteases, and neutrophils release neutrophil extracellular traps (NETs), which exacerbate tissue damage [7] [8] [9] [10] .

Several studies on SARS-CoV2 infection have attempted to elucidate phenotypic features of immune cell subsets either associated with severity or predictive of disease outcome. However, these studies have been conducted, in most cases, with a limited number of patients, and clinical parameters and disease severity are not homogeneously recorded in all of them [11, 12] . Likewise, although specific antibodies and complement activation have been proposed to mediate some of the most severe complications of coronavirus infections, including that of SARS-CoV-2 [2, 5, 13, 14] , solid evidence on the role of humoral immunity effector mechanisms in the pathogenesis of SARS-CoV-2-associated ARDS is clearly needed.

Our cohort of COVID-19 patients included 276 SARS-CoV-2 infected individuals who were further classified according to the severity of their clinical signs and symptoms in mild, moderate and severe, following recently described criteria [15] . The mean duration of symptoms before admission was 7.36 ± 5.2 days. Table 1 shows their main demographic and laboratory characteristics. The median (percentile 25 and 75) age was 63 (53.25-75) and 163 (59.05%) were men.

We conducted an initial analysis following the COVID-19 admission protocol, which comprised the quantification of the main peripheral blood lymphocyte subsets, including T, B and NK lymphocytes as well as plasmablasts, by multi-parametric flow cytometry (Fig.1A, Suppl. Fig 1) . The proportion of T lymphocytes (either CD4+ or CD8+) and NK cells was similar in all the patients and healthy volunteers, with the exception of CD8+ T cells, which decreased in severe patients (Fig.1B) .

Conversely, the percentage of B cells was higher in COVID-19 patients and increased with disease severity, raising from a mean of 9.15% in healthy donors to 20.49% in severely ill patients. In accordance, plasmablasts were remarkably higher in patients, both in relative and absolute values (mean 1.51% vs 17.98%; and 2.83 cells/l vs 27.37 cells/l, respectively), and regardless severity degree ( Figure 1B ). This increase in absolute plasmablasts number is of special relevance, given the lymphopenia present in COVID-19 patients (Table 1) , and the diminished absolute number of other lymphocyte subsets (Fig.1B) . The elevated number of plasmablasts suggested a redistribution of the main maturation stages of the B lineage, including naïve, transitional, unswitched memory, IgM-only memory and class-switched memory B-cells, which were identified in a subgroup of 84 patients from our initial cohort, with the gating strategy shown in Suppl. Fig 2. In COVID-19 patients, the proportion of IgM-only memory B-cells increased, while unswitched memory cells decreased (Fig.1C) . This redistribution was more marked in mild patients and progressively diminished in moderate and severe patients. Nevertheless, differences among patients with different severity degree were not statistically significant (Suppl. Fig 3) .

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Given the role of Tfh in maturation and activation of B cells, we assessed whether circulating Tfh (cTfh) were increased in the peripheral blood of COVID-19 patients, in accordance with the increased number of plasmablasts. We observed that the cTfh proportion significantly increased with the severity of COVID-19 individuals, shifting from a median of 0.51% in healthy donors to 1.7 % in severely ill patients. Importantly, the absolute number remained steady despite the profound decrease of total CD4+ T cells ( Fig.2A, Fig.1B) . To further characterize this population, we assessed surface expression of CCR7 chemokine receptor, which has been related to lower B-cell activation capacity by Tfh [16, 17] . We found a significant increase of CCR7 expression in cTfh cells of patients with moderate to severe disease (Fig 2A) . Finally, we found a direct correlation between cTfh proportion and total B-cells (r 0.33; p= 0.0090), class-switched B-cells (r 0.26, p= 0.0433), and plasmablasts (r 0.33, p= 0.0091) in peripheral blood (Fig.2B ).

The high number of plasmablasts prompted us to investigate possible alterations in Ig concentrations. At the time of admission, COVID-19 patients had serum concentrations of either IgG, IgA or IgM isotypes comparable to those in healthy volunteers (Fig.3A) . On the other hand, when patients with different degree of severity were analysed, we observed that, in severe cases, the levels of IgA and IgM were similar, but the IgG concentration was decreased, compared to healthy volunteers (Fig.3B ). In addition, a direct correlation was found between IgG (0.37; p=0.0007) and IgA The increase in plasma concentrations of IL-6 and acute phase reactants that characterizes COVID-19 (Table 1) suggested that the concentration of C3 and C4 complement proteins, considered as acute phase reactants, could also be elevated. Therefore, we measured C3 and C4 levels in the sera of these patients, and could detect increased levels of both complement proteins (Fig.3A) . Accordingly, to investigate whether complement activation occurs in COVID-19 patients, we quantified C5a, the activation peptide of complement component C5, which showed increased plasma levels in most patients ( Figure 3A ).

However, when considering different groups of severity, we observed that C3 and C4 levels increased in patients with mild to moderate disease, while returned to levels similar to healthy donors in those with severe disease (Fig.3B) . Therefore, in contrast to other inflammatory parameters such as LDH, ferritin or CRP (Table 1) , C3 and C4 values decreased as the severity increased. Of note, unlike C3 and C4, C5a levels remained elevated in plasma regardless degree of severity (Fig.3B ).

Next, we tested whether the decrease in these components of humoral immunity was actually related to the severity of the disease in critical patients, or rather mirrored an increased consumption along time. We then considered a hospitalization period longer than 15 days as a readout of the severity of the disease and collected the blood of a subgroup of 37 COVID-19 patients who were still hospitalized 10 days after admission and compared it to the initial blood test. With this approach, we observed that antibodies of the IgG, IgA and IgM isotypes, the IgG subclasses IgG1, IgG3, and IgG4 as well as C3, C4 and C5a complement proteins were similar in all patients at the time of admission ( Fig.4A and B) . However, after 10 days, serum concentration of IgG was significantly lower in patients hospitalized longer than 15 days, whereas levels of IgA and IgM did not change over this time. Interestingly, whereas IgG1 and IgG3 levels showed a significant fold decrease in long vs. short stay patients after 10 days from admission, IgG4 concentration diminished similarly in both groups of patients. In order to determine whether these changes corresponded to a specific anti-SARS-CoV-2 response, we quantified IgG against SARS-CoV-2 RBD and N proteins, which could be detected at a high titer at day 10 in all cases (Suppl. Fig.4 ). On the contrary, anti-CMV IgG levels decreased after 10 days of hospitalization in both groups (Suppl. Fig 4) .

Regarding complement, C3 serum levels were stable over time in both groups of patients. On the other hand, a significant decrease of C4 concentration was observed after 10 days specifically in those patients whose severity eventually required a longer stay (Fig.4A ). These patients however showed sustained C5a levels at this time point (Fig.4A) . Conversely, activation product C5a decreased in patients who stayed less than 15 days, which did not show a parallel C4 significant reduction (Fig.4A ).

The decrease in C4 and IgG serum concentrations in severe COVID-19 cases led us to hypothesize that antigen-antibody complexes were forming in excessive amounts and activating the classical pathway of complement system. Therefore, we quantified immune complexes by enzyme immunoassay in the serum of the SARS-Cov-2 infected individuals with different levels of severity, but we did not find detectable levels (data not shown).

The decrease of IgG that characterized severe patients also suggested a possible role of ADCC, which has been described to be mediated by specific NK cells subsets [18] . We therefore quantified the main functional subsets of NK cells (CD56 bright CD16-, CD56 dim CD16+, CD56-CD16+). A two-fold increase in the proportion of CD56-CD16+ NK cells was found in severe COVID-19 patients compared This article is protected by copyright. All rights reserved.

to healthy donors and patients with mild disease (Fig. 5A) . Furthermore, the absolute values of this population remained similar to those of healthy donors in all COVID-19 patients, in contrast to the decrease seen for other lymphocyte populations (Fig. 5A ). Further analysis of this population showed a significant downregulation of CD16 expression irrespective of the degree of severity (Fig.5B) .

Interestingly, CD16 level in CD56-CD16+ cells slightly correlated to the serum concentration of IgG in COVID-19 patients (r-0.24; p=0.0496) (Fig.2B ).

It has been proposed that the severity of COVID-19 is related to a dysregulation of the immune Plasmablasts were significantly elevated in peripheral blood of most COVID-19 patients at the time of admission. In contrast, Ig levels were similar to those of healthy donors, or even decreased in the case of IgG in critically ill patients. Since patients were studied at admission, it was feasible that Ig production had not peaked yet. However, IgG levels further decreased 10 days after admission in patients with longer hospital stays, which suggests that IgG decrease was rather related to severe disease progression. Three possible explanations may account for these apparently contradictory observations. The first explanation is a primary antibody immunodeficiency that debuts with COVID-19. This is supported by the fact that several patients had very low levels of IgG, IgA and IgM in serum both at admission and after 10 days. We have not addressed this possibility yet, however we plan to carry out studies aimed at verifying this hypothesis by assessing Ig levels three months after discharge. Second, it is possible that certain individuals show impaired specific Ig production (as might appear from the decrease of the titer of CMV-specific antibodies shown in supplementary figure 4) after infection secondary to generalized lymphopenia, altered B cell maturation, or patient age. Regarding impaired specific Ig production, it is known that sepsis triggers immune cell hyperactivity followed, days later, by immune paralysis, which associates with a poor patient The nature of specific antibodies which could activate complement should, therefore, require further research. For example, a recent study in a SARS-CoV macaque model that appeared before the emergence of first cases of COVID-19, demonstrated that a faster development of neutralizing IgG against SARS-CoV spike characterized animals who developed severe lung injury [14] . Moreover, S glycoprotein-specific Ab responses were higher and peaked earlier in deceased patients during the first 15 days after the appearance of symptoms, but dramatically dropped 5 days later, coinciding with clinical deterioration [30] .

In our cohort, we have found that IgG1 and IgG3 isotypes, but not IgG4, decreased after 10 days from admission specifically in more severe patients, a finding that could be related to the greater capacity of those isotypes to mediate the activation of the classical pathway of complement. Of note, specific IgG3 antibodies are the first to appear in viral infections [31] . Likewise, C3 and C4 levels were increased in both mild and moderate patients but not in individuals with severe COVID-19, where C3 and C4 remained within the normal values or were even lower. The reasons for these findings include individual variability in C3 and C4 synthesis due to genetic deficiencies or individual genetic polymorphisms [32] [33] [34] . Alternatively, a higher C3 and C4 consumption may occur in severe patients by an excessive activation of either the lectins pathway or the classical pathway through specific anti-viral antibodies or immune-complexes. The selective C4 depletion observed is probably due to the limited amount of C4 in plasma compared to C3, which is one of the most abundant plasma proteins. In addition, a clear correlation between C3 and C4 was found in those patients (data not shown), suggesting a parallel consumption of both proteins which are characteristic of some autoimmune diseases and severe infections [35] [36] [37] [38] . C5a is one of the strongest proinflammatory molecules, since it has chemoattractant activity and activates most leukocytes. In particular, C5a promotes the lung sequestration of myeloid cells [40] and pulmonary dysfunction and exerts procoagulant activity through several mechanisms, including the induction of tissue factor by endothelial cells and neutrophils, as well as the upregulation of Thus, while immune system is required for protection from SARS-CoV-2 infection, its response, if excessive or sustained over time, like that occurring in COVID-19, may perpetuate both ongoing inflammation and the hypercoagulable state. It is therefore necessary to provide more insights related to individual characteristics and the particular kinetics of the host response to SARS-CoV-2, to further understand the pathogenesis of COVID-19 and to decide the appropriate therapy and the treatment timing. This knowledge is of particular therapeutic relevance since it is the basis for clinical trials with the different immune modulators that are currently available including intravenous immunoglobulins, hyperimmune plasma or complement inhibitors, among others [59] [60] [61] .

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This is a retrospective observational study including 276 consecutive patients with confirmed detection of SARS-CoV-2 RNA, and admitted to the Accident and Emergency Department of the Hospital Universitario La Princesa because of mild to critical COVID-19 symptoms, from February 27 th to April 29 th .

Demographic and laboratory data described in Table 1 were collected from electronic clinical records and included in an anonymized database. Baseline evaluation of immunity was performed around the 3 rd day of admission (median=3 days; percentile 25-75 [p25-p75] 2 to 6). A second evaluation was obtained around the 14th day of admission (median=14 days; p25-p75 12 to 16.5) in a selected group of 37 patients.

Samples from nasopharyngeal and throat exudates were obtained with specific swabs as previously described [62] . As first line screening, we performed real-time RT-PCR assay targeting the E gene of SARS-CoV-2, with Real Time ready RNA Virus Master on Applied Biosystems TM Quant Studio-5 Real-Time PCR System. This assay was followed by confirmatory testing with the assay TaqPath™ COVID-19 CE-IVD Kit RT-PCR Applied Biosystems™ (ThermoFisher Scientific, Waltham, MA USA), which contains a set of TaqMan RT PCR assays for in vitro diagnostic use. This kit includes three assays that target SARS-CoV-2 genes (Orf1ab, S gene, N gene) and one positive control assay that targets the human RNase P RPPH1 gene [63] . Determinations were carried out in an Applied Biosystems TM QuantStudio-5 Real-Time PCR System (CA, USA). 

Nucleoprotein (NP) and the Receptor Binding Domain (RBD) of SARS-CoV-2 were produced and ELISA Serum samples were also assessed with an anti-CMV/IgG detection kit (Enzygnos, Siemens), following manufacturer's instructions.

Descriptive results were expressed as mean ± standard deviation (SD) or median and percentile 25percentile 75 (p25-p75), as appropriate, while qualitative variables are presented as frequency (n) and relative percentages of patients (%). The unpaired, two-tailed, Student t-test was used to compare two independent groups and the paired Student t-test, to analyse two related samples.

One-way ANOVA was employed to compare more than two groups and post-hoc multiple comparisons were made with Tukey's test. Spearman bivariate correlations were performed between serological quantitative markers and cell populations and corrplot R package (available from: https://github.com/taiyun/corrplot) was used for correlation map graphics. Variables in correlation map were reordered using hierarchical cluster method. The p-values were two-sided and statistical significance was considered when p < 0.05. To analyse distribution of lymphocyte in COVID-19 patients and healthy donors, an automated clustering and dimensionality reduction was performed using viSNE and FlowSOM tools (Cytobank). Population cells were normalized using log transformation for analysis. Differences in normalized cells between healthy donors and severity groups (adjusted by sex and age) were assessed with a moderated t-test using limma R package [66] .

Cell populations that showed significant P-value (FDR = 5%) were considered as differentially expressed between groups. Stata v. 12.0 for Windows and R version 3.5.1 were used for analyses and graphics. GraphPad Prism 4 software was also used for graphics. Data is presented making specification for p < 0.05 (*), p < 0.01 (**), p < 0.001 (***) and p < 0.0001 (****).

This study was approved by the local Research Ethics Committee (register number 4070) and it was carried out following the ethical principles established in the Declaration of Helsinki. All included patients were informed about the study and gave an oral informed consent because of COVID-19 emergency as proposed by AEMPS (Agencia Española del Medicamento y Productos Sanitarios). Table I . Demographic and laboratory characteristics of the study population classified by severity degree. All variables are expressed as median (p25-p75). AST: aspartate amino-transferase; ALT: alanine amino-transferase; GGT: gamma-glutamyl transferase; LDH: lactate dehydrogenase; CRP: Creactive protein; IL-6: interleukin-6. . Asterisks indicate significant differences (p-values for ANOVA Tukey's contrast test: *p < 0.05, **p < 0.01, ***p < 0.001, **** p < 0.0001) (B) Annotated heatmap of a correlation matrix for different variables in those same COVID-19 patients (n= 84). White squares include non-significant correlations (p > 0.05), red and blue squares include significant indirect and direct correlations (p < 0.05), respectively. Numbers inside squares and intensity of color correspond to Spearman's rank correlation coefficient. Variables in the correlation map were reordered using hierarchical cluster method. Data are from 19 experiments, with a median of 6 patients per experiment and day. Boxplots display percentiles 25 and 75 and median, and the whiskers correspond to percentiles 5 and 95. and COVID-19 patients (n=63). (B) Serum concentration (mg/dL) of IgG, IgA, IgM, C3 and C4 in healthy donors (n=19) and COVID-19 patients according to severity degree (mild (n= 138), moderate (n= 82) and severe (n= 35). Serum concentration (ng/mL) of C5a in healthy donors (n=10) and COVID-19 patients according to severity degree (mild (n= 29), moderate (n= 19) and severe (n= 15). Values represent quantification for each serum marker depicted as boxplots. Asterisks indicate significant differences (p-values for Mann-Whitney t-test or ANOVA Kruskal-Wallis test, as appropriate: *p < 0.05, **p < 0.01, ***p < 0.001, **** p < 0.0001). Data are from samples collected over a 6 weeks period and analyzed within 24 hours before freezing the serum. For C5a measurements, samples were thawed and assessed in 2 independent experiments. Boxplots depict percentiles 25 and 75 and the median, and the whiskers show percentiles 5 and 95. between day 0 and day +10 after admission. Green boxplots correspond to patients with a hospitalization period of less than 15 days (n= 13) and orange boxplots to those hospitalized more than 15 days (n= 24). C5a quantification in orange boxplots corresponds to n=15. Asterisks indicate significant differences (p-values for Mann-Whitney t-test or Wilcoxon t-test, as appropriate: *p < 0.05, **p < 0.01, ***p < 0.001, **** p < 0.0001). Data are from samples collected over a 6 weeks period and analyzed within 24 hours before freezing the serum. For IgG1, IgG3 and IgG4 levels measurements, samples were thawed and assessed within 2 independent experiments.. Boxplots display percentiles 25 and 75 and the median, and the whiskers correspond to percentiles 5 and 95. Boxplot depict CD16 MFI of CD56 dim CD16+ and CD56-CD16+ NK subpopulations in those same individuals. Asterisks indicate significant differences (p-values for ANOVA Tukey's contrast test: *p < 0.05, **p < 0.01, ***p < 0.001, **** p < 0.0001). Data are from 19 experiments, with a median of 6 patients per experiment and day. Boxplots depict percentiles 25 and 75 and the median, and the whiskers correspond to percentiles 5 and 95.

Graphical abstract text for "Deregulated cellular circuits driving immunoglobulins and complement consumption associate with the severity of COVID-19 patients"

The pathogenesis of SARS-CoV-2 involves both cellular and humoral immunity. The proportion of B cells, plasmablasts, circulating follicular helper T cells (cTfh) and CD56-CD16+ NK-cells is increased in COVID-19 patients. However, levels of IgM, IgG and complement proteins are diminished in severe patients, this suggesting immunoglobulins and complement consumption.

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The study was funded by grants SAF2017-82886-R to FS-M from the Ministerio de Economía y Competitividad, and from "La Caixa Banking Foundation" (HR17-00016) to FS-M. Grant PI018/01163 to CMC and grant PI19/00549 to AA were funded by Fondo de Investigaciones Sanitarias, Ministerio de Sanidad y Consumo, Spain. SAF2017-82886-R, PI018/01163 and PI19/00549 grants were also cofunded by European Regional Development Fund, ERDF/FEDER. This work has been funded by grants Fondo Supera COVID (CRUE-Banco de Santander) to FSM, and "Ayuda Covid 2019" from Comunidad de Madrid.We thank Dr. Miguel Vicente-Manzanares for proofreading and English editing of the manuscript.We also thank the immunology service staff for technical support: Victor López-Huete, Alicia Román, Reyes Lázaro-Tejedor, Alicia Vara-Vega, Montserrat Arroyo-Correa, Manuela Mayo. The Graphical Abstract was created with BioRender.com.

The authors declare no commercial or financial conflict of interest. This article is protected by copyright. All rights reserved.

The data that support the findings of this study are available from the corresponding author upon reasonable request.