Abstract
Despite extensive and ongoing studies of SARS-CoV-2 and evidence that pregnant women are at increased risk of severe COVID-19, the effect of maternal infection on the developing infant remains unclear. To determine the potential impact of exposure to SARS-CoV-2 in utero on the neonate, we have assessed the immunological status of infants born to mothers with confirmed SARS-CoV-2 infection during gestation. No evidence of vertical transmission of SARS-CoV-2 was observed, but transfer of maternal SARS-CoV-2 specific IgG to infants was apparent, although to a lesser extent in cases of active or recent maternal infection. Infants born to mothers with recent/ongoing infection had elevated circulating pro-inflammatory cytokines and enhanced percentages of innate immune cells compared to that seen in infants born to uninfected mothers. In tandem, higher frequencies of FOXP3+ regulatory T cells and circulating IL-10 demonstrated a further nuance to the neonatal effector response. Interestingly, cytokine functionality was enhanced in infants born to mothers exposed to SARS-CoV-2 at any time during pregnancy. This indicates that maternal SARS-CoV-2 infection influences in utero priming of the fetal immune system.
Introduction
Despite the ongoing COVID-19 pandemic, the effect of maternal SARS-CoV-2 infection on the immunology of the developing infant is still unclear. Indeed, SARS-CoV-2 infection in pregnancy has been reported to lead to variable outcomes for the mother. The majority of infected women in pregnancy are asymptomatic or only experience mild symptoms 1,2. Nevertheless, some pregnant women with SARS-CoV-2, especially in the third trimester, appear to be at an increased risk for hospitalization and subsequent intensive care unit admission 3,4,5 and rates of maternal infection increased in the second wave 6,7. Although the rates of preterm birth did not appear elevated in initial reports, more recent data suggest pregnant women are at a higher risk for subsequent preterm birth albeit much may be due to clinical intervention 8.
With respect to the infant, a national UK surveillance study suggested neonatal SARS-CoV-2 infection is uncommon even in babies born to mothers with perinatal infection 9. Similarly, a study from the US of 116 mothers with confirmed perinatal SARS-CoV-2 infection did not identify any neonatal cases 10 and a systematic review demonstrated no vertical transmission 11. Set against this, there have been a small number of individual case reports documenting evidence of vertical transmission12,13,14,15,16,17, although it was subsequently established that only 30% of documented cases were due to true placental transfer 18.
Whilst vertical transmission of SARS-CoV-2 itself is rare, the potential immunological perturbations induced in the pregnant mother 19,20 may conceivably leave an immunological legacy on the newborn with far reaching consequences. Indeed, our recent study in a preterm baby group found evidence of perinatal inflammation modulating the developing immune system of the infant 21. It is already appreciated that the immune system of the unborn child can be altered by the presence of human immunodeficiency virus (HIV) or Hepatitis C virus (HCV) in mothers, either with or without vertical transmission 22,23,24,25; that metabolites derived from maternal gut microbiota can shape the immune system of the offspring 26; and that modulation of the neonatal immune system has been associated with diseases in later life 27,28. In the limited studies that have assessed the immune status of babies born to SARS-CoV-2 infected mothers, there has been little evidence of impact in the cellular and humoral immunity of the neonate 29. Similarly, a small study of SARS-CoV-2 infection during pregnancy has been associated with a cytokine response in the fetal circulation (i.e. umbilical cord blood) but with no effect on the cellular immune repertoire 20. However, to our knowledge, none of the studies have included a comprehensive analysis of the infant cellular immune profile of babies born to SARS-CoV-2 exposed mothers (at any point in their pregnancy) compared to profiles of infants born to unexposed mothers. Indeed, it is increasingly challenging to include an appropriate control group in a global pandemic. Moreover, simultaneous analysis of cytokines and antibody titres in infants and their paired mothers is lacking. In our study, we have assessed the immunologic status of infants born to mothers with SARS-CoV-2 that tested positive either during the two weeks directly prior to birth or earlier on in pregnancy, compared to babies born to mothers never exposed to SARS-CoV-2 to identify if there is a legacy of maternal infection and potential in utero priming of the neonatal immune system.
Results
Transfer of passive immunity is reduced in infants born to mothers with recent/ongoing infection
Maternal and infant characteristics are shown in Table 1, and the group description is shown in Fig. 1a. Our neonatal group of infants born to mothers with SARS-CoV-2 exposure (SARS-CoV-2 exposed, SE, n=30) was divided into those with recent or ongoing infection as determined by mothers confirmed positive for SARS-CoV-2 by PCR within 2 weeks of birth (R/O, n=16, median positive swab 3 days before birth), or prior to 2 weeks; termed the recovered group (R, n=14, median positive swab 48.5 days prior to birth).
In both the recent/ongoing and recovered group, regardless of the time between the positive swab and birth or maternal IgM levels, SARS-CoV-2 specific IgM was not detected in cord plasma, suggestive of a lack of vertical transmission (Fig. 1b). This was true for IgM directed against the spike protein (S), the receptor binding domain within the spike protein (RBD) as well as the nucleoprotein (N) for which background IgM reactivity has been shown to be higher 30. By contrast, SARS-CoV-2 specific IgG against all three antigens was detected in infants born to SARS-CoV-2 exposed mothers (Fig. 1b). Whilst levels of SARS-CoV-2 specific IgG in the mother-infant dyad were comparable in the recovered group, there were significantly lower levels of SARS-CoV-2 specific IgG in infants born to mothers with recent/ongoing infection (Extended Data Fig. 1a). Thus, when the ratio of infant Ig to their paired maternal Ig (transfer ratio) was calculated for each antigen, the mean transfer ratio of all 3 antigens was 1.04 in the R group and 0.79 in the R/O group (Fig. 1c). This was despite the presence of high levels of maternal IgG in at least some mothers in the R/O group (Extended Data Fig. 1a). Indeed, when only comparing IgG levels in infants born to seropositive mothers, transfer of SARS-CoV-2 specific IgG to the infant was still significantly lower in the R/O group (Fig. 1d) and this did not appear to differ with the sex of the infant (Extended Data Fig. 1b).
Elevated plasma cytokines in mothers with recent/ongoing infection and their infants
SARS-CoV-2 infection is known to be associated with marked elevation of several plasma cytokines including Interferon gamma-induced protein 10 (IP-10), interleukin (IL)-1β, CXCL8, IL-6 and IL-10 31,32,33,34. To assess the impact of this on the neonate, plasma cytokine concentrations were assessed in paired maternal and cord blood using a multiplex assay. IP-10 and IL-1β levels in the plasma of mothers with recent/ongoing SARS-CoV-2 infection were significantly elevated when compared to that of recovered mothers whilst IL-10, CXCL8 and IL-6 were similar between the SE groups (Fig. 2a). When assessing neonatal cytokine levels, IL-10 was significantly elevated in the cord plasma from babies born to mothers with recent/ongoing infection compared to those born to recovered mothers. CXCL8 levels were also numerically higher in the recent/ongoing group although this did not reach significance, largely driven by three infants with undetectably levels (Fig. 2b). However, concentrations of this chemokine was significantly higher in infants than in their paired mothers (Fig. 2c) which was not seen with any of the other cytokines tested. Conversely, infant IP-10 was significantly lower than their paired mothers (Extended Data Fig. 2a). The majority of babies born to recovered mothers that showed elevated levels of CXCL8 were born by vaginal delivery (Fig. 2d), known to elevate several cytokines 35. However, there were still notable increases of CXCL8 in babies born via caesarean section (CS) in the recent/ongoing group, compared to the recovered group, suggesting this was indeed a fetal response to maternal SARS-CoV-2 infection.
Spearman rank test identified significant (p<0.05) correlations in cytokine levels both within and between maternal and cord blood in both groups. Nonetheless, this was more evident in mothers and infants in the R/O group compared with the recovered group suggestive of a greater degree of immune co-regulation in recent/ongoing SARS-CoV-2 infection (Fig. 2e): there was a correlation between CXCL8 and IP-10 in infants born to mothers with recent/ongoing infection (Fig. 2f); maternal IL-1β levels correlated with CXCL8 and IP-10 in the infants from this group (Fig. 2g-h), and interestingly, maternal IL-1β levels negatively correlated with days between a positive COVID swab and birth (Fig. 2i), suggesting this cytokine was indicative of recent infection in the mothers. No differences were seen in plasma IL-12p70, GM-CSF, IFN-α2, IFN-λ1, IFN-λ2/3, IFN-β, TNF-α and IFN-γ in mothers or infants from either group (Extended Data Fig. 2b).
Recent/ongoing maternal SARS-CoV-2 infection influences adaptive immune cell populations in the infant
To determine the potential impact of maternal SARS-CoV-2 infection on the cellular immune compartment of the neonate at birth, we employed multiparametric flow cytometry to phenotype peripheral blood leukocytes and assess their in vitro functional capacity upon mitogen stimulation (Figs. 3-5). Gating strategies are shown in Extended Data Fig. 3. To establish if maternal SARS-CoV-2 infection altered the developing immune system of the neonate, infant cellular immune profiles in the combined (R and R/O) SARS-CoV-2 exposed group (SE; n=30) were compared to those from term infants born to healthy mothers collected prior to the pandemic (Non SARS-CoV-2 exposed, NSE; n=15) but measured contemporaneously. We performed tSNE dimensionality reduction (Fig. 3a) on 91 individual flow cytometry immune parameters and observed that infant immune profiles in the SE group clustered away from the normal immune profiles of the NSE group. Correlations between immune parameters appeared significantly different in infants born to SE mothers compared to NSE mothers (Fig. 3b-c). For example, a weaker correlation was seen between IFN-γ producing cells and a stronger correlation occurred between proliferating TEMRAs (Extended Data Fig. 4a-b). Indeed, when focusing on the immune parameters that drive the biggest differences, we identified that infants born to mothers in the R/O group separated furthest away from the NSE group, using 3-dimensonal PCA (Fig. 3d), and that all three groups tended to segregate based on their maternal SARS-CoV-2 status upon unbiased hierarchal clustering analysis (Fig. 3e). Moreover, there was no clear clustering based on alternative confounding factors such as sex, ethnicity, mode of delivery and other maternal characteristics such as chorioamnionitis and gestational diabetes (GDM) (Extended Data Fig. 5). Despite the segregation observed in the infant cellular immune compartment, many parameters, including those known to be perturbed in the adult response to SARS-CoV-2, were not different between groups. Infant T cell lymphopenia was not observed and moreover the relative frequencies of major adaptive lymphocyte subsets (e.g. CD4 and CD8 αβ T cells, γδ T cells and B cells) were unaffected by maternal exposure to SARS-CoV-2 (Fig. 3f). Despite preserved composition of cell frequencies, changes in adaptive cell populations were still evident, including an increase in the percentage of CD161 expressing CD8 T cells in babies born to mothers with recent/ongoing infection (Fig. 3g) as well as increased CD25+FOXP3+ TREGS (Fig. 3h), the latter of which positively correlated with the percentage of Vδ2 T cells (Fig. 3i).
Conspicuous increased proportions of innate immune cells in neonates born to mothers with recent/ongoing SARS-CoV-2 infection
In contrast to the adaptive immune cell compartment, changes in innate cell subsets were more prominent. NK, NKT and innate-like Vδ2 γδ T cells were significantly elevated in babies born to mothers with recent/ongoing infection (Fig. 4a-c). There was also a change in monocyte populations with enhanced percentages of alternative monocytes and subsequently reduced CD38+ classical monocytes in babies born to mothers with recent/ongoing infection (Extended Data Fig. 6a). These cells were, for the most part, not significantly elevated in infants born to mothers with previous SARS-CoV-2 exposure (R), consistent with reactive neonatal responses to recent/ongoing SARS-CoV-2 infection. Indeed, raised percentages of NK cells negatively correlated with days from positive SARS-CoV-2 swab result to birth, further suggesting this was a neonatal response to maternal infection (Extended Data Fig. 6b). Interestingly, the percentage of NKT cells (Fig. 4d) and NK cell activation (as assessed by CD69 expression, Fig. 4e) both positively correlated with the levels of cord blood CXCL8 suggesting these are key immune markers associated with the neonatal response to maternal infection.
Increased cytokine functionality in innate and adaptive cells in all infants born to SARS-CoV-2 exposed mothers
When assessing functionality by intracellular cytokine staining post polyclonal activation, the ability of immune cells to produce cytokines upon stimulation was significantly elevated in babies born to SE mothers. Consistent with changes observed in plasma cytokine concentrations and cellular immune composition, enhanced cytokine production by neonatal immune cells was associated with maternal SARS-CoV-2 infection. Indeed, cytokine potential was significantly enhanced in several different cell types in babies born to mothers previously exposed to SARS-CoV-2 (at any time point) as exemplified by TNF-α (Fig 5a), IFN-γ (Fig. 5b) and to a lesser extent, IL-17 (Extended Data Fig. 6c) although the ability to produce CXCL8 was unaltered (Extended Data Fig. 6d). The observed enhanced cytokine functionality (TNF-α and IFN-γ) in CD4 T cells positively correlated with effector memory CD4 T cells (Fig. 5 c,d), and TNF-α producing cells (both CD4 and CD8) negatively correlated with CD38 expression, known to decrease during maturation (Extended Data Fig. 6e) 21. Due to limited numbers of mothers with severe disease, we were unable to establish if the extent of immune imprinting was related to maternal infection status.
Discussion
Our study provides a comprehensive immune atlas of neonates born to mothers with SARS-CoV-2 exposure. Whilst we did not observe vertical transmission of SARS-CoV-2 itself, we did find multiple immunological perturbations within the neonate associated with maternal SARS-CoV-2 exposure during pregnancy, many of which were also associated with recent/ongoing infections. Taken together, our findings are suggestive of an immunological legacy imprinted on the neonate following maternal SARS-CoV-2 exposure, with potential far-reaching consequences. In utero exposure to environmental factors, infection and/or maternal inflammation is increasingly recognised to affect the developing immune system and subsequent responses both to infection 36, immune mediated diseases 37-39 and neurodevelopmental problems 40,41. Indeed, long term effects cannot be ruled out as observed in survivors after in utero exposure to the 1918 (Spanish) influenza pandemic 42,43.
Although we did not directly assess neonates for the presence of SARS-CoV-2, we did assess SARS-CoV-2 specific IgM levels and could find no evidence of vertical transmission in any of the 30 infants born to SARS-CoV-2 exposed mothers. SARS-CoV-2 specific IgG was, however, transferred to the neonates from their mothers suggestive of the transfer of protective immunity. There was a correlation between maternal and infant SARS-CoV-2 IgG levels in mother-infant dyads in both groups as previously suggested 44. However, even though for many pathogens, umbilical cord titers of IgG at normal term delivery are higher than in maternal blood 45,46,47,48, there were reduced levels of SARS-CoV-2 specific IgG in infants born to mothers with recent/ongoing infection compared to their paired mothers. This did not appear to be a threshold issue, as many mothers exhibited high levels of SARS-CoV-2 specific IgG which was not transferred efficiently to their infant. Reduction of SARS-CoV-2 specific Ig transfer via the placenta has been suggested to occur in the third trimester due to altered glycosylation 49 and reduced maternal SARS-CoV-2-specific antibody titers and impaired placental antibody transfer were also noted in pregnancies with a male fetus 50, although there did not appear to be any sex bias in this data set. It is currently unclear whether antibodies induced via vaccination as opposed to natural infection, differ in terms of their glycosylation status and subsequent placental transfer. Vaccination to SARS-CoV-2 in the second and third trimester did elicit placental transfer of Abs, with a reduced transfer ratio observed in the last trimester51,52.. Our study adds further evidence suggesting the 2nd trimester may represent more opportune vaccination timing, at least with respect to the transfer of passive immunity to the infant.
Perhaps unsurprisingly, cord plasma of neonates born to mothers with recent/ongoing infection, expressed elevated concentrations of some cytokines known to be associated with inflammation and COVID-19, consistent with placental immune activation 31. Previously, elevated levels of IP-10, IL-6, IL-10, CXCL8 and IL-1β have been associated with adult infection and the former three with disease severity 32,33,34. Increased IL-6 and IL-10 have also been associated with severity in early SARS-CoV-2 infection in children 53. Interestingly, this conventional COVID-19 signature of adults was skewed more towards IL-10 (and to a lesser extent, CXCL8) in the neonates born to mothers with recent/ongoing infection. Theoretically, the increased concentrations of cytokines in the infant could be explained by transfer of maternal cytokines through the placental tissues. Indeed, infants with raised IP-10 concentrations were born to mothers who had high plasma IP-10 measurements, although in general concentrations were significantly lower in infants compared to mothers. However, in the case of cord plasma CXCL8, a chemokine which correlated strongly with maternal IL-1β (previously associated with maternal SARS-CoV-2 infection, 54) levels were significantly higher than that observed in their mothers, suggesting that at least some of these elevated cytokines were fetal-derived, and a direct response to maternal infection. Elevated cord plasma cytokines, including CXCL8, have been observed in some infants born to SARS-CoV-2+ mothers 20. Similarly, increased cord plasma CXCL8 has been observed after in utero exposure to HIV infection 55, and in pregnant women with GBS infection and/or chorioamnionitis 56,57, suggesting that maternal infection can directly influence fetal CXCL8. Raised fetal inflammatory markers, therefore, do appear to be related to the maternal infection status at the time of birth, as such profiles are less obvious in infants born to mothers with recovered infection. Indeed, where we did see slight elevations in CXCL8 (and IL-6) in the recovered group, this could be explained by the mode of delivery, as labour is known to drive elevation of these cytokines 35.
The well-documented lymphopenia observed in COVID-19 and also in children with MIS-C 58,59 was not observed in infants born to mothers either with SARS-CoV-2 infection around birth or mothers who had previously had infection. This may be partially attributable to the proposed mechanisms for the observed peripheral lymphopenia, including sequestration of lymphocytes to affected organs such as the lung, direct viral invasion of T cells via ACE2 binding, or virus-induced destruction of secondary lymphoid organs 60 none of which apply to the infant in the absence of viral transmission. Indeed, in contrast to COVID-19 infection where reductions in peripheral NK, NKT, Vδ2 and MAIT cells have been observed in adults 32,61,62,63, neonates born to mothers exposed to SARS-CoV-2 actually exhibited elevated percentages of NK, NKT and Vδ2 γδ T cells and also CD161 expressing CD8 T cells (the majority of which are likely to be MAIT cells). These innate-like cells may be responding to the inflammatory cytokine milieu in the context of maternal infection potentially as a protective response in the neonate vis-à-vis their likely beneficial role in severe adult COVID-19 62. Immune activation of these cells has been seen at the maternal fetal interface 31 and although a previous report suggested there was no elevation of NK cells in infants born to COVID-19 mothers 29, this was only compared to reported reference levels 64 and there was no direct comparator group in their study.
As well as alterations in these cell populations, we also identified enhanced cytokine potential upon in vitro stimulation. This was observed not only in infants born to mothers with recent/ongoing infection but also in those born to recovered mothers which suggests potential in utero priming of the immune response. Hence, the percentage of CD4, CD8, NK, NKT or γδ T cells that produced TNF-α (or IFN-γ and IL-17 to a lesser extent) was significantly greater in infants born to mothers exposed to SARS-CoV-2. At birth neonatal T cells predominantly produce CXCL8 with a limited capacity for IFN-γ and IL-17 production that increases with age 65). Thus, these findings may reflect some accelerated maturation of the neonatal immune system induced in utero by maternal SARS-CoV-2 infection. Much of this enhanced cytokine functionality correlated with other markers of immune maturation such as increased percentages of memory T cells and decreased T cell expression of CD38. IFN-γ expression is controlled by epigenetic mechanisms in neonates 66, so it is tempting to speculate that maternal SARS-CoV-2 infection may have induced some epigenetic changes in these loci. Indeed, maternal exposure to polycyclic aromatic hydrocarbons directly altered this locus in cord blood mononuclear cells 67 and other data suggests that epigenetic modifications during gestation can shape the future development of diseases like obesity, type 2 diabetes, allergy, asthma, and infections 68,69. Increased proportions of cytokine producing T cells have also been observed following in utero exposure to malaria, HBV and HIV-even without vertical transmission 70-72.
Taken together, these data strongly suggest that maternal SARS-CoV-2 infection shapes the immune profile of an infant to different extents dependent on the time of exposure. We identified a transient response to maternal inflammation in the form of increased cytokines in cord plasma but also altered immune cell functionality in neonates exposed to SARS-CoV-2 at any point during gestation, suggesting some immune imprinting. Whilst the aetiology of the observed immune perturbations in the neonate remain unclear, the consequences could be far-reaching.
More data are needed to establish if these changes specifically relate to enhanced protection from SARS-CoV-2 mediated disease or are detrimental if the infant is born in a milieu of inflammatory cytokines. A hyper-inflammatory MIS-C like response has been observed in a neonate following in utero exposure to SARS-CoV-2 (mother infected 9 weeks prior to birth) with no evidence of direct neonatal infection 73. Long term follow-up of the infants in our study will establish if maternal exposure to SARS-CoV-2 has a long lasting impact on the child. These data may also have implications regarding the vaccination regimen for pregnant women. Indeed, the reduced transfer of protective Abs to the infant we observed in those infants born to mothers with recent infection may suggest 2nd trimester or early 3rd trimester vaccination is preferable. Neonatal immune profiling following vaccination in pregnancy may also determine what level of maternal immune activation drives the neonatal imprinting observed.
Methods
Study design and human material
Umbilical cord blood (and paired maternal peripheral blood) was collected over the COVID-19 pandemic (28th May 2020-1st March 2021) at the time of birth from infants born to mothers who were SARS-CoV-2 Exposed (SE) attending the maternity unit at GSTT, London in heparinised blood tubes at the time of birth to investigate the immune status (REC Approval No. 19/SC/0232). Mother infant dyads were categorised into recent/ongoing (R/O) and recovered groups (R), according to the number of days prior to birth that the mother received a positive status from a SARS-CoV-2 nasopharyngeal swab (R/O: <14 days; R: ≥14 days). Umbilical cord blood was also collected at the time of birth from infants born to healthy mothers at GSTT prior to the COVID-19 pandemic (until 1st Jan 2020), hence their mothers were not infected with SARS-CoV-2 at any time during their pregnancy, termed the Non SARS-CoV-2 Exposed (NSE) group (REC Approval No. 17/LO/0641). The clinical details of the groups can be found in Table 1.
Isolation of CMBCs and plasma
Cord blood mononuclear cells (CBMCs) were isolated from all infant groups. Neat plasma was also isolated from the recent/ongoing and recovered groups and their paired mothers (all SE samples). More specifically, for the paired maternal peripheral blood samples, whole blood was centrifuged for 2000 g for 10 min at room temperature, and plasma was harvested from the upper layer and stored at -80°C in polypropylene tubes. For the neonatal samples, cord blood was layered onto Ficoll (GE Healthcare), within a 15 ml polypropylene conical tube, and centrifuged at 800 g for 15 min (break OFF) at room temperature. Plasma was then collected from the top fraction and stored at - 80°C in polypropylene tubes. The CBMC layer was isolated and subsequently washed twice with pre-warmed base medium (BM) (RPMI-1640 + L-Glutamine, Gibco), and then complete medium (CM) (RPMI-1640 + L-Glutamine; 10% Heat-Inactivated FBS; 1% Penicillin-Streptomycin, Gibco), under centrifugation at 300 g for 5 min at room temperature. The cell pellet was then frozen in Cryostor® CS10 (Sigma) within polypropylene cryovials before proceeding with flow cytometry.
Polyclonal stimulation
Prior to staining the CBMCs for flow cytometry immunophenotyping with panel 4 (Extended Data Table 1), cells were thawed and plated in 96 well round bottom plates (Corning) within 200 μL CM containing Phorbol 12-myristate 13-acetate (PMA) (10 ng/ml) (Sigma), Ionomycin (1 μg/ml) (Sigma), Brefeldin A (20 ng/ml) (Sigma) and Monensin solution (1x) (BioLegend). A Brefeldin A and Monensin only control was also plated for each infant. Cells were incubated at 37°C for 4 h in the Biosafety level 3 (BSL-3) containment lab, in accordance with the King’s College London safety rules before proceeding with flow cytometry staining. Cells were removed from BSL-3 once they were fixed with Cell Fix (1X) (BD) for a minimum of 10 mins.
Flow cytometry staining and acquisition
CBMCs were thawed and plated in 96 well round bottom plates before staining in one of 4 panels (Extended Data Table 1) assessing the following immune cell populations (panel 1: T cell naïve/memory status; panel 2: myeloid and B cells; panel 3: T and NK cell activation status; panel 4: T and NK cytokine potential). All four panels contained surface marker staining, and panel 1 and 4 also contained intracellular staining. All the following wash steps were performed under 2000 g, for 1 min at room temperature. For each panel, cells were washed with 100 μL Dulbecco’s Phosphate Buffered Saline (1X) (PBS, Gibco) and resuspended in 100 μL PBS, containing Zombie NIR™ Fixable Viability dye (1:1000) (Biolegend), with the addition of TCR Vδ1-FITC (TS8.2) (Thermo Fisher) in panel 1, for 15 mins in the dark at 4°C. Cells were then washed with 150 μL eBioscience™ FOXP3 Fixation/Permeabilization buffer (Invitrogen), for panel 1, or FACS buffer (PBS, 0.5% heat-inactivated FBS, 2mM EDTA, Invitrogen), for panels 2-4. The wash step was repeated with 200 μL volume. Cells were then resuspended in 100 μL eBioscience™ FOXP3 Permeabilization buffer (FPB) (Invitrogen), for panel 1, or 50 μL surface antibody cocktail within FACS buffer, for panels 2-4, for 30 mins (panels 2-3) or 20 mins (panel 4), at 4°C in the dark. Cells were subsequently washed in 100 μL FPB (panel 1) or 150 μL FACS buffer (panels 2-4), and again with 200 μL volume before resuspending in 50 μL antibody cocktail in FPB (panel 1), or 100 μL Cell Fix (1x) (panels 2-4) for 30 mins (panels 1-3) or until intracellular cytokine staining (ICS) (panel 4) at 4°C in the dark. Panels 1-3 were then washed twice in 100 μL FPB (panel 1) or FACS buffer (panels 2-3) and resuspended in 200 μL FACS buffer until acquisition. For the panel 4 ICS, cells were centrifuged at 2000 g, 1 min, at room temperature and resuspended in 50 μL ICS antibody cocktail within Permeabilization Wash Buffer (1x) (PWB) (BioLegend) for 30 mins in the dark at room temperature. Cells were washed twice in 150 μL PWB, followed by 200 μL FACS buffer before resuspending the cells in FACS buffer for acquisition in a high-throughput sampler on a 4-laser LSR Fortessa (BD), at a flow rate of 1 μL/s.
Flow cytometry data analysis
Raw FCS files were analysed using FlowJo (v10.6.2, BD) and the gating strategies are included in Extended Data Fig. 3. As an internal control, the same adult sample was run alongside each flow cytometry experiment for consistency and to aid setting gates. Data were cleaned up by gating on the Time parameter to ensure that only cells going through a constant flow stream were analysed, and cell populations were excluded from downstream analysis if the event count in the parent population was <30.
SARS-CoV-2 specific antibody testing
ELISAs were conducted as previously described 30,74. All plasma samples were heat-inactivated at 56 °C for 30 min before use. High-binding ELISA plates (Corning, 3690) were coated with antigen (Nuclear protein (N), Spike glycoprotein (S) or the receptor binding domain (RBD) at 3 µg ml-1 (25 µl per well) in PBS, either overnight at 4 °C or for 2 h at 37 °C. Wells were washed with PBS-T (PBS with 0.05% Tween-20) and then blocked with 100 µl of 5% milk in PBS-T for 1 h at room temperature. Wells were emptied and serial dilutions of plasma (starting at 1:25, 5-fold dilution) were added and incubated for 2 h at room temperature. Control reagents included CR3009 (2 µg/ml) (N-specific monoclonal antibody), CR3022 (0.2 µg/ml) (S-specific monoclonal antibody), negative control plasma (1:25 dilution), positive control plasma (1:50), and blank wells. Wells were washed with PBS-T. Secondary antibody was added and incubated for 1h at room temperature. IgG was detected using goat-anti-human-Fc-AP (1:1000) (Jackson, catalogue no. 109-055-098) and wells were washed with PBS-T and AP substrate (Sigma) was added and plates read at 405 nm. IgM was detected using goat-anti-human-IgM-HRP (1:1000) (Sigma catalogue no. A6907) and wells were washed with PBS-T and one-step 3,3′,5,5′-tetramethylbenzidine (TMB) substrate (Thermo Fisher Scientific) was added and quenched with 2M H2S04 before reading at 450 nm (HRP). Samples were run in duplicate and peak fold-change above background was plotted for each patient. IgG transfer ratios were calculated to display the difference in paired infant and maternal levels [Infant IgG peak fold change over background/maternal IgG peak fold change over background].
Quantification of plasma cytokines
Paired maternal and infant plasma were thawed and tested in the 13-plex LegendPlex Human Anti-Virus Response Panel kit (Biolegend), to quantify levels of IL-1β, IL-6, TNF-α, IP-10, CXCL8, IL-12p70, IFN-α2, IFN-λ1, IFN-λ2/3, GM-CSF, IFN-β, IL-10 and IFN-γ. The assay was performed according to the manufacturer instructions and was modified by diluting the kit reagents 2x (beads, detection antibodies and streptavidin-PE). All plasma samples were diluted 2x with assay buffer, and resulting sample concentrations were calculated according to the dilution factor. In brief, 25 μL diluted plasma or standard, and mixed beads (1:1 ratio) were added to each well (in V-bottom 96-well plates) and incubated for 1.5 h. The samples were washed twice with wash buffer, incubated with 25 μL detection antibodies for 1 h and then 25 μL streptavidin-PE was added for a further 30 mins. The samples were then washed once with wash buffer, resuspended in 200 μL wash buffer and acquired on a high-throughput sampler with a 3-laser FACSCanto™ (BD). All incubation steps were performed under 600 rpm on an orbital shaker at room temperature and protected from light. Data were cleaned up by excluding cytokines if the bead event count was <90, to ensure accurate analyses, performed using the Windows LegendPlex (v8.0, BioLegend) software.
Statistical analyses
Analysed flow cytometry populations, plasma cytokines and antibodies were imported into an Excel spreadsheet and analysed in R (v4.0.3) to generate boxplots, dimensionality reduction plots, Spearman correlation plots and heatmaps. Clustered heatmaps were performed on scaled and centred data using the heatmap.2 package, and clustered according to the Euclidean method. The corrplot package was used to generate Spearman correlation matrices, and only significant values (p<0.05) are displayed. GraphPad Prism (v9.0) was also utilised to generate scatter plots for cytokines and the antibody heatmap. All statistical tests were used to measure the differences between biologically distinct samples. Unadjusted p values (*p<0.05; **p<0.01; ***P<0.001 and ****p<0.0001) were assessed by the Kolmogorov-Smirnov test (to compare cytokine concentrations between the groups), two-sided Wilcoxon rank-sum tests (for immune cell populations between the groups), and two-sided paired Wilcoxon tests (between paired maternal and infant antibody/cytokine levels).
Author Contributions
MC and CM: Patient consent, study design and sample collection. SG: Sample processing, flow cytometry and data analysis. SG and AD: panel design, multiplex cytokine testing and analysis. JS and KJD: Antibody ELISA testing and analysis. DG and RT: Manuscript writing and experimental design. All authors reviewed drafts of the manuscript prior to submission.
Competing Interests
The authors declare no competing interests.
Acknowledgements
We thank the mothers and their infants for blood collection and all the midwives at STH for sample collection. We also thank Shraddha Kamdar and those staff involved with COVID-IP at King’s College London 32 for some sample processing and Thomas Lechmere for assistance with ELISAs. We thank Iva Zlatareva for assistance with the cytokine multiplex analysis. We would also like to acknowledge Evolve Biosystems for funding (RT and DG) towards the non SARS-CoV-2 exposed (NSE) infant cord blood sample collection and the staff involved (Niamh Kelly, Lucy McMillan, Sarah Kheirallah and Jiadai Mi). We thank Yin Wu for critical reading of the manuscript. SG is supported by a MRC-KCL Doctoral Training Partnership in Biomedical Sciences (MR/N013700/1), DLG by Action Medical Research (GN2790), MC is supported by the NIHR BRC COVID-19 call, RMT by Tommy’s (Charity No. 1060508) and Borne (1167073) and JS and KJD by Kings Together Rapid COVID-19 call award and Huo Family Foundation award. The research was also supported by the National Institute for Health Research (NIHR) Biomedical Research Centre based at Guy’s and St Thomas’ NHS Foundation Trust (GSTT) and King’s College London (KCL) (part of the King’s Health Partners Academic Sciences Centre). The views expressed are those of the authors and not necessarily those of the NHS, the NIHR or the Department of Health.