Pregnant women are at increased risk of adverse outcomes, including preeclampsia and preterm birth, that may result from severe acute respiratory syndrome coronavirus 2 (SARS-CoV-2) infection. Pregnancy imprints specific maternal immune responses that can modulate host susceptibility to microbial infection; therefore, recent studies have focused on the humoral response against SARS-CoV-2 in pregnant women. However, the pregnancy-specific cellular immune responses triggered by SARS-CoV-2 infection are poorly understood. In this study, we undertook an extensive in vitro investigation to determine the cellular immune responses to SARS-CoV-2 particles and proteins/peptides in pregnant women. First, we show that SARS-CoV-2 particles do not alter the pregnancy-specific oxidative burst of neutrophils and monocytes. Yet, SARS-CoV-2 particles/proteins shift monocyte activation from the classical to intermediate states in pregnant, but not in nonpregnant, women. Furthermore, SARS-CoV-2 proteins, but not particles or peptide pools, mildly enhance T cell activation during pregnancy. As expected, B cell phenotypes are heavily modulated by SARS-CoV-2 particles in all women; yet, pregnancy itself further modified such responses in these adaptive immune cells. Lastly, we report that pregnancy itself governs cytokine responses in the maternal circulation, of which IFN-β and IL-8 were diminished upon SARS-CoV-2 challenge. Collectively, these findings highlight the differential in vitro responses to SARS-CoV-2 in pregnant and nonpregnant women and shed light on the immune mechanisms implicated in coronavirus disease 2019 during pregnancy.

The coronavirus disease 2019 (COVID-19) pandemic caused by the severe acute respiratory syndrome coronavirus 2 (SARS-CoV-2) (1) is still an ongoing global catastrophe (2, 3). Nearly 450 million people have been infected, and >6 million worldwide have died (2). Notably, women are more likely to exhibit reduced COVID-19 severity and mortality compared with men (4, 5). However, pregnant women are not included in such a disparity, given their increased risk of adverse outcomes associated with COVID-19, including admission to the intensive care unit, need for mechanical ventilation, and death (68). Furthermore, pregnant women with COVID-19 are more likely to undergo obstetrical complications (911), as evidenced by a correlation between the severity of SARS-CoV-2 infection in pregnancy and the risk of preeclampsia (9, 10), preterm birth (10), and stillbirth (11). Yet, SARS-CoV-2 is not the only virus to which pregnant women are more susceptible (7, 12). Increased risk of adverse outcomes in pregnant women has also been reported for other viruses, such as influenza, SARS-CoV-1, and Middle East respiratory syndrome coronavirus (1315). Importantly, neonates born to women exposed to viruses during pregnancy can subsequently develop chronic diseases during adulthood, as evidenced by the increased prevalence of cardiovascular disease in adults born to women affected by the Spanish flu in 1918 (16, 17). Therefore, elucidating the underlying mechanisms leading to such heightened susceptibility during pregnancy is timely.

Pregnancy is a unique immunological state in which the maternal immune system undergoes complex and tightly regulated adaptations to allow the mother to tolerate the semiallogeneic fetus (1821) and vice versa (2224). This is contrary to the historical view that pregnancy is an immunosuppressive state (2527). Indeed, the innate immune limb of pregnant women is strengthened in its capacity to deal with viral or bacterial infection, showing an increase in the number of granulocytes and oxidative burst as well as altered phenotypes in basal and stimulated conditions (2830). By contrast, pregnant women display decreased numbers of T and B cells in the peripheral blood (30), yet they exhibit enhanced immune cellular components of innate and adaptive origin such as homeostatic macrophages (31, 32) and regulatory T cells (3336). Hence, pregnancy itself imprints specific maternal immune responses (37), which can dictate susceptibility or resistance to microbes. Notably, recent studies have characterized the humoral (i.e., Ab-mediated) response toward SARS-CoV-2 Ags in pregnant and nonpregnant women (3840). However, the pregnancy-specific cellular immune responses implicated in SARS-CoV-2 infection are poorly understood.

In the current study, we performed an extensive in vitro investigation to evaluate the cellular immune responses to SARS-CoV-2 particles and proteins/peptides in pregnant women and compared such to nonpregnant women. First, we explored whether SARS-CoV-2 particles alter reactive oxygen species (ROS) production by neutrophils and monocytes as a readout of the acute innate host response. Next, monocyte phenotypes and activation status in response to SARS-CoV-2 particles and proteins/peptides were characterized by using in vitro assays coupled with flow cytometry. Furthermore, T cell activation, as evidenced by the expression of surface markers and effector molecules, and B cell phenotypes were also evaluated in response to SARS-CoV-2 particles and proteins/peptides. Finally, by using multiplex immunoassays, cytokine release was determined in PBMCs and in whole-blood TruCulture systems in response to SARS-CoV-2 particles and spike (S) protein, respectively.

Peripheral blood samples were collected from healthy nonpregnant and pregnant women. All women were recruited into research protocols of the Perinatology Research Branch, an intramural program of the Eunice Kennedy Shriver National Institute of Child Health and Human Development, National Institutes of Health, U.S. Department of Health and Human Services, Wayne State University (Detroit, MI), and the Detroit Medical Center (Detroit, MI). The collection and use of human materials for research purposes were approved by the Institutional Review Boards of Wayne State University School of Medicine and the Detroit Medical Center. All participating women provided written informed consent prior to sample collection. The maternal peripheral blood of pregnant women was collected in the third trimester prior to the administration of any medication. The nonpregnant group included healthy women of reproductive age from the same community as the pregnant patients. Women who self-reported as testing positive for SARS-CoV-2, having COVID-19, or receiving a vaccine were excluded from the study.

Peripheral blood samples were collected from individuals by venipuncture into EDTA-containing tubes. After collection, 50 µl of whole blood were stimulated with 50 µl of ROS assay mix containing a 1:250 dilution of ROS assay stain in ROS assay buffer (ROS Assay Kit; catalog no. 88-5930; eBioscience, San Diego, CA) together with 1 µl of PMA (3 µg/ml; catalog no. P1585-1MG; Millipore Sigma-Aldrich, Burlington, MA) or 1 µl of 1× PBS (catalog no. 10-010-023; Thermo Fisher Scientific, Durham, NC) as an unstimulated control. The cells were incubated at 37°C with 5% CO2 for 60 min. Following incubation, erythrocytes were lysed using ammonium-chloride-potassium lysing buffer (Quality Biological, Gaithersburg, MD), and the resulting leukocytes were collected after centrifugation at 300 × g for 5 min. Leukocytes were resuspended in 0.5 ml of 1× PBS and acquired with the BD LSRFortessa flow cytometer (BD Biosciences, San Jose, CA) and FACSDiva 9.0 software (BD Biosciences) to measure ROS production by neutrophils and monocytes. Data analysis and plots were performed with FlowJo software version 10 (Tree Star, Ashland, OR).

PBMCs were isolated from blood samples collected in EDTA-containing tubes by using the density-gradient reagent Lymphoprep (catalog no. 07801; StemCell Technologies, Vancouver, British Columbia, Canada), according to the manufacturer’s instructions. Mononuclear cell suspensions were cultivated in complete RPMI 1640 medium (catalog no. 11875-093; Thermo Fisher Scientific, Life Technologies Limited, Paisley, U.K.) (enriched with 5% human serum [catalog no. H3667; Sigma-Aldrich]) and 1% penicillin-streptomycin (catalog no. 15140122; Thermo Fisher Scientific) at 1 × 106 cells/ml in a six-well culture plate. Heat-inactivated SARS-CoV-2 culture fluid (41) (containing viral particles and hereafter termed “SARS-CoV-2 particles”) (Isolate USA-WA1/2020; catalog no. 0810587CFHI; ZeptoMetrix Corporation, Buffalo, NY) was added to the PBMC culture at 1 × 106 50% tissue culture infectious dose/ml, and culture supernatants from Vero E6 cells alone were added as controls. Cells were incubated at 37°C and 5% CO2 for 48 h with the addition of a protein transport inhibitor mixture (catalog no. 00-4980-93; Thermo Fisher Scientific) for the last 4 h of incubation. Adherent cells were gently lifted from the plate and collected. Cell suspensions were transferred into 5-ml Falcon tubes and centrifuged at 300 × g for 5 min at 4°C. Cell supernatants were harvested and stored at −80°C for cytokine determinations, and cell pellets were used for immunophenotyping.

PBMCs were isolated as described above and cultivated in complete RPMI 1640 medium in a 12-well culture plate at 1 × 106 cells/ml for monocyte stimulation or in a 24-well culture plate at 2.5 × 106 cells/ml for T cell stimulation. SARS-CoV-2 nucleocapsid (N) protein (catalog no. 40588-V08B), SARS-CoV-2 S protein (S protein receptor binding domain; catalog no. 40592-V08B), SARS-CoV-2 S protein (S1 Subunit; catalog no. 40591-V08B1), SARS-CoV-2 S protein (S1+S2 extracellular domain; catalog no. 40589-V08B1), and SARS-CoV-2 (2019-nCoV) S1 (S1 protein mutant D614G; catalog no. 40591-V08H3) (all from Sino Biological, Wayne, PA) were added to their respective cell culture well at final concentrations of 20 nM. A nonprotein control, a positive control for T cell activation (PHA; 5 μg/ml; catalog no. L8902; Sigma-Aldrich), and a positive control for monocyte activation (LPS of Escherichia coli, 100 ng/ml; catalog no. tlrl-peklps; Invivogen, San Diego, CA) were also included. Plates were incubated at 37°C and 5% CO2 for 24 h (for monocytes) or 48 h (for T cells). Protein transport inhibitor mixture was added to the wells for the last 4 h of incubation. Cells were then collected, transferred into 5-ml Falcon tubes, and centrifuged at 300 × g for 5 min at 4°C. Cell supernatants were stored at −80°C for cytokine determinations, and cell pellets were used for immunophenotyping.

PBMCs were isolated as described above and cultivated in complete RPMI 1640 medium in a 12-well culture plate at 1 × 106 cells/ml for monocyte stimulation or in a 24-well culture plate at 2.5 × 106 cells/ml for T cell stimulation. PepTivator SARS-CoV-2 Prot_S (catalog no. 130-126-700), PepTivator SARS-CoV-2 Prot_S1 (receptor binding domain) (catalog no. 130-127-041), PepTivator SARS-CoV-2 Prot_N (catalog no. 130-126-698), PepTivator SARS-CoV-2 Prot_M (catalog no. 130-126-702), and PepTivator CEF MHC Class I Plus–premium grade (catalog no. 130-098-426; peptide-specific positive control) (all from Miltenyi Biotec, Bergisch Gladbach, Germany) were added to their respective cell culture well at final concentrations of 0.6 nmol/ml. A nonpeptide control, a positive control for T cell activation (PHA; 5 μg/ml), and a positive control for monocyte activation (LPS; 100 ng/ml) were also included. Plates were incubated at 37°C and 5% CO2 for 24 h for monocytes and 48 h for T cells. Protein transport inhibitor mixture was added to the cell cultures for the last 4 h of incubation. Cells were then collected, transferred into 5-ml Falcon tubes, and centrifuged at 300 × g for 5 min at 4°C. Cell supernatants were stored at −80°C for cytokine determinations, and cell pellets were used for immunophenotyping.

After stimulation, PBMC pellets were washed and resuspended in 1× PBS followed by incubation with Fixable Viability Stain 510 (catalog no. 564406; BD Biosciences) for 15 min at room temperature in the dark. Then, the cells were washed and subsequently incubated with specific extracellular fluorochrome-conjugated anti-human mAbs (Table I) for 30 min at 4°C in the dark. After washing, the cells were fixed and permeabilized with either the BD Cytofix/Cytoperm kit (catalog no. 554714; BD Biosciences) or the eBioscience Foxp3/Transcription Factor Staining Buffer Set (catalog no. 00-5523-00; Thermo Fisher Scientific, Life Technologies Corporation, Carlsbad, CA) prior to staining with intracellular Abs (Table I). Leukocytes were then washed and resuspended in 0.5 ml of FACS staining buffer (catalog no. 554656; BD Biosciences) and acquired with the BD LSRFortessa flow cytometer and FACSDiva 9.0 software. Data analysis and plots were performed with the FlowJo software version 10. Immunophenotyping included the identification of general leukocyte populations (monocytes, T cells, and B cells), monocyte subsets, T cell subsets, and B cell subsets. Specifically, classical monocytes (CD14hiCD16), intermediate monocytes (CD14hiCD16+ cells), nonclassical monocytes (CD14loCD16+), and CD14loCD16 cells were identified as well as CD4+ and CD8+ T cells, CD19+ B cells, and their respective memory or activation markers.

Table I.

List of Abs used for immunophenotyping

AgFluorochromeCloneCompanyIsotype
CD14 BUV395 MφP9 BD Biosciences, catalog no. 563561 Mouse BALB/c IgG2b, κ 
CD16 APC-H7 3G8 BD Biosciences, catalog no. 560195 Mouse CDF1 IgG1, κ 
CCR5 (CD195) BV711 2D7/CCR5 BD Biosciences, catalog no. 563395 Mouse C57BL/6 IgG2a, κ 
CD181 (CXCR1) PE-Cy5 5A12 BD Biosciences, catalog no. 551081 Mouse IgG2b, κ 
CD182 (CXCR2) APC 6C6 BD Biosciences, catalog no. 551127 Mouse IgG1, λ 
CX3CR1 PE-Cy7 2A9-1 Biolegend, catalog no. 341612 Rat IgG2b, κ 
CD142 (TF) BUV737 HTF-1 BD Biosciences, catalog no. 748835 Mouse IgG1, κ 
CD147 (Basigin) PerCP-Cy5.5 HIM6 BD Biosciences, catalog no. 562554 Mouse IgG1, κ 
CD162 (PSGL) BV786 KPL-1 BD Biosciences, catalog no. 743483 Mouse BALB/c IgG1, κ 
CCR2 BV650 LS132.1D9 BD Biosciences, catalog no. 747849 Mouse BALB/c IgG2a, κ 
IL-6 PE-CF594 MQ2-13A5 BD Biosciences, catalog no. 563543 Rat IgG1 
IL-8 BV421 G265-8 BD Biosciences, catalog no. 563310 Mouse IgG2b 
TNF BV605 MAb11 BioLegend, catalog no. 502936 Mouse IgG1, κ 
CXCL10 Alexa Fluor 488 4NY8UN Invitrogen, catalog no. 53-9744-42 Mouse/IgG2b, κ 
IL-1RA PE AS17 BD Biosciences, catalog no. 340525 Mouse IgG1 
CD3 PerCP-Cy5.5 UCHT1 BD Biosciences, catalog no. 560835 Mouse BALB/c IgG1, κ 
CD4 BUV737 SK3 BD Biosciences, catalog no. 612748 Mouse BALB/c IgG1, κ 
CD8 BUV395 RPA-T8 BD Biosciences, catalog no. 563795 Mouse IgG1, κ 
CD25 PE-Cy7 M-A251 BD Biosciences, catalog no. 557741 Mouse BALB/c IgG1, κ 
CD45RA Alexa Fluor 700 HI100 BD Biosciences, catalog no. 560673 Mouse IgG2b, κ 
CCR7 PE 3D12 BD Biosciences, catalog no. 552176 Rat IgG2a, κ 
CD154 (CD40L) FITC TRAP1 BD Biosciences, catalog no. 558988 Mouse BALB/c IgG1, κ 
CD137 (4-1BB) BV421 4B4-1 BD Biosciences, catalog no. 564091 Mouse BALB/c IgG1, κ 
CD69 PE-Cy5 FN50 BD Biosciences, catalog no. 555532 Mouse IgG1, κ 
CD134 (OX40) APC-Cy7 Ber-ACT35 BioLegend, catalog no. 350022 Mouse IgG1, κ 
CD95 (FAS) APC DX2 BD Biosciences, catalog no. 558814 Mouse C3H/Bi IgG1, κ 
Granzyme B PE-CF594 GB11 BD Biosciences, catalog no. 562462 Mouse BALB/c IgG1, κ 
IFN-γ BV650 4S.B3 BD Biosciences, catalog no. 563416 Mouse BALB/c IgG1, κ 
IL-10 BV786 JES3-9D7 BD Biosciences, catalog no. 564049 Rat IgG1 
Ki67 BV711 B56 BD Biosciences, catalog no. 563755 Mouse IgG1, κ 
CD19 Alexa Fluor 488 HIB19 BD Biosciences, catalog no. 557697 Mouse IgG1, κ 
CD20 PerCP-Cy5.5 2H7 BD Biosciences, catalog no. 560736 Mouse C57BL/6 IgG2b, κ 
CD23 Alexa Fluor 700 M-L233 BD Biosciences, catalog no. 563673 Mouse IgG1, κ 
CD27 APC-H7 M-T271 BD Biosciences, catalog no. 560222 Mouse BALB/c IgG1, κ 
CD86 PE-CF594 2331(FUN-1) BD Biosciences, catalog no. 562390 Mouse BALB/c IgG1, κ 
CD80 BV650 2D10.4 BD Biosciences, catalog no. 751725 Mouse IgG1, κ 
CD40 PE-Cy7 5C3 BD Biosciences, catalog no. 561215 Mouse IgG1, κ 
CD43 BV605 1G10 BD Biosciences, catalog no. 563378 Mouse IgG1, κ 
CD22 PE-Cy5 HIB22 BD Biosciences, catalog no. 555425 Mouse IgG1, κ 
CD24 BUV395 ML5 BD Biosciences, catalog no. 563818 Mouse IgG2a, κ 
CD38 BUV737 HB7 BD Biosciences, catalog no. 612824 Mouse IgG1, κ 
CD138 BV711 MI15 BD Biosciences, catalog no. 563184 Mouse BALB/c IgG1, κ 
CD5 BV421 UCHT2 BD Biosciences, catalog no. 562646 Mouse BALB/c IgG1, κ 
IgD PE IA6-2 BD Biosciences, catalog no. 555779 Mouse BALB/c IgG2a, κ 
IgM APC G20-127 BD Biosciences, catalog no. 551062 Mouse IgG1, κ 
IgG BV786 G18-145 BD Biosciences, catalog no. 564230 Mouse IgG1, κ 
Isotype BV711 X40 BD Biosciences, catalog no. 563044 Mouse BALB/c IgG1, κ 
Isotype PE-Cy5 MOPC-21 BD Biosciences, catalog no. 555750 Mouse IgG1, κ 
Isotype APC MOPC-21 BD Biosciences, catalog no. 555751 Mouse IgG1, κ 
Isotype PE-CF594 R3-34 BD Biosciences, catalog no. 562309 Rat IgG1, κ 
Isotype BV421 27-35 BD Biosciences, catalog no. 562748 Mouse C.SW IgG2b, κ 
Isotype BV605 MOPC-21 BioLegend, catalog no. 400162 Mouse IgG1, κ 
Isotype Alexa Fluor 488 eBMG2b Invitrogen, catalog no. 53-4732-80 Mouse/IgG2b, κ 
Isotype PE MOPC-21 BD Biosciences, catalog no. 555749 Mouse IgG1, κ 
Isotype PE-CF594 X40 BD Biosciences, catalog no. 562292 Mouse BALB/c IgG1, κ 
Isotype BV650 X40 BD Biosciences, catalog no. 563231 Mouse BALB/c IgG1, κ 
Isotype BV786 R3-34 BD Biosciences, catalog no. 563847 Rat IgG1, κ 
AgFluorochromeCloneCompanyIsotype
CD14 BUV395 MφP9 BD Biosciences, catalog no. 563561 Mouse BALB/c IgG2b, κ 
CD16 APC-H7 3G8 BD Biosciences, catalog no. 560195 Mouse CDF1 IgG1, κ 
CCR5 (CD195) BV711 2D7/CCR5 BD Biosciences, catalog no. 563395 Mouse C57BL/6 IgG2a, κ 
CD181 (CXCR1) PE-Cy5 5A12 BD Biosciences, catalog no. 551081 Mouse IgG2b, κ 
CD182 (CXCR2) APC 6C6 BD Biosciences, catalog no. 551127 Mouse IgG1, λ 
CX3CR1 PE-Cy7 2A9-1 Biolegend, catalog no. 341612 Rat IgG2b, κ 
CD142 (TF) BUV737 HTF-1 BD Biosciences, catalog no. 748835 Mouse IgG1, κ 
CD147 (Basigin) PerCP-Cy5.5 HIM6 BD Biosciences, catalog no. 562554 Mouse IgG1, κ 
CD162 (PSGL) BV786 KPL-1 BD Biosciences, catalog no. 743483 Mouse BALB/c IgG1, κ 
CCR2 BV650 LS132.1D9 BD Biosciences, catalog no. 747849 Mouse BALB/c IgG2a, κ 
IL-6 PE-CF594 MQ2-13A5 BD Biosciences, catalog no. 563543 Rat IgG1 
IL-8 BV421 G265-8 BD Biosciences, catalog no. 563310 Mouse IgG2b 
TNF BV605 MAb11 BioLegend, catalog no. 502936 Mouse IgG1, κ 
CXCL10 Alexa Fluor 488 4NY8UN Invitrogen, catalog no. 53-9744-42 Mouse/IgG2b, κ 
IL-1RA PE AS17 BD Biosciences, catalog no. 340525 Mouse IgG1 
CD3 PerCP-Cy5.5 UCHT1 BD Biosciences, catalog no. 560835 Mouse BALB/c IgG1, κ 
CD4 BUV737 SK3 BD Biosciences, catalog no. 612748 Mouse BALB/c IgG1, κ 
CD8 BUV395 RPA-T8 BD Biosciences, catalog no. 563795 Mouse IgG1, κ 
CD25 PE-Cy7 M-A251 BD Biosciences, catalog no. 557741 Mouse BALB/c IgG1, κ 
CD45RA Alexa Fluor 700 HI100 BD Biosciences, catalog no. 560673 Mouse IgG2b, κ 
CCR7 PE 3D12 BD Biosciences, catalog no. 552176 Rat IgG2a, κ 
CD154 (CD40L) FITC TRAP1 BD Biosciences, catalog no. 558988 Mouse BALB/c IgG1, κ 
CD137 (4-1BB) BV421 4B4-1 BD Biosciences, catalog no. 564091 Mouse BALB/c IgG1, κ 
CD69 PE-Cy5 FN50 BD Biosciences, catalog no. 555532 Mouse IgG1, κ 
CD134 (OX40) APC-Cy7 Ber-ACT35 BioLegend, catalog no. 350022 Mouse IgG1, κ 
CD95 (FAS) APC DX2 BD Biosciences, catalog no. 558814 Mouse C3H/Bi IgG1, κ 
Granzyme B PE-CF594 GB11 BD Biosciences, catalog no. 562462 Mouse BALB/c IgG1, κ 
IFN-γ BV650 4S.B3 BD Biosciences, catalog no. 563416 Mouse BALB/c IgG1, κ 
IL-10 BV786 JES3-9D7 BD Biosciences, catalog no. 564049 Rat IgG1 
Ki67 BV711 B56 BD Biosciences, catalog no. 563755 Mouse IgG1, κ 
CD19 Alexa Fluor 488 HIB19 BD Biosciences, catalog no. 557697 Mouse IgG1, κ 
CD20 PerCP-Cy5.5 2H7 BD Biosciences, catalog no. 560736 Mouse C57BL/6 IgG2b, κ 
CD23 Alexa Fluor 700 M-L233 BD Biosciences, catalog no. 563673 Mouse IgG1, κ 
CD27 APC-H7 M-T271 BD Biosciences, catalog no. 560222 Mouse BALB/c IgG1, κ 
CD86 PE-CF594 2331(FUN-1) BD Biosciences, catalog no. 562390 Mouse BALB/c IgG1, κ 
CD80 BV650 2D10.4 BD Biosciences, catalog no. 751725 Mouse IgG1, κ 
CD40 PE-Cy7 5C3 BD Biosciences, catalog no. 561215 Mouse IgG1, κ 
CD43 BV605 1G10 BD Biosciences, catalog no. 563378 Mouse IgG1, κ 
CD22 PE-Cy5 HIB22 BD Biosciences, catalog no. 555425 Mouse IgG1, κ 
CD24 BUV395 ML5 BD Biosciences, catalog no. 563818 Mouse IgG2a, κ 
CD38 BUV737 HB7 BD Biosciences, catalog no. 612824 Mouse IgG1, κ 
CD138 BV711 MI15 BD Biosciences, catalog no. 563184 Mouse BALB/c IgG1, κ 
CD5 BV421 UCHT2 BD Biosciences, catalog no. 562646 Mouse BALB/c IgG1, κ 
IgD PE IA6-2 BD Biosciences, catalog no. 555779 Mouse BALB/c IgG2a, κ 
IgM APC G20-127 BD Biosciences, catalog no. 551062 Mouse IgG1, κ 
IgG BV786 G18-145 BD Biosciences, catalog no. 564230 Mouse IgG1, κ 
Isotype BV711 X40 BD Biosciences, catalog no. 563044 Mouse BALB/c IgG1, κ 
Isotype PE-Cy5 MOPC-21 BD Biosciences, catalog no. 555750 Mouse IgG1, κ 
Isotype APC MOPC-21 BD Biosciences, catalog no. 555751 Mouse IgG1, κ 
Isotype PE-CF594 R3-34 BD Biosciences, catalog no. 562309 Rat IgG1, κ 
Isotype BV421 27-35 BD Biosciences, catalog no. 562748 Mouse C.SW IgG2b, κ 
Isotype BV605 MOPC-21 BioLegend, catalog no. 400162 Mouse IgG1, κ 
Isotype Alexa Fluor 488 eBMG2b Invitrogen, catalog no. 53-4732-80 Mouse/IgG2b, κ 
Isotype PE MOPC-21 BD Biosciences, catalog no. 555749 Mouse IgG1, κ 
Isotype PE-CF594 X40 BD Biosciences, catalog no. 562292 Mouse BALB/c IgG1, κ 
Isotype BV650 X40 BD Biosciences, catalog no. 563231 Mouse BALB/c IgG1, κ 
Isotype BV786 R3-34 BD Biosciences, catalog no. 563847 Rat IgG1, κ 

Peripheral blood samples were collected from women by venipuncture into heparin-containing tubes. One milliliter of whole blood was directly placed into the TruCulture (Myriad RBM, Austin, TX) (42) tubes containing SARS-CoV-2 S protein (part no. 782-001411) or null tubes (part no. 782-001086) as a control. The TruCulture tubes were gently inverted 10 times to mix the whole blood and culture media and then incubated for 48 h at 37°C in a heating block. After incubation, cell supernatants were collected separately into cryovials and stored at −80°C.

Cell culture supernatants and TruCulture tube supernatants were utilized to determine cytokine concentrations. The V-PLEX Proinflammatory Panel 1 (human) and U-PLEX Biomarker Group 1 (human) multiplex immunoassays (Meso Scale Discovery, Rockville, MD) were used to measure the concentrations of IFN-γ, IL-1β, IL-2, IL-4, IL-6, IL-8, IL-10, IL-12p70, IL-13, and TNF (Proinflammatory Panel 1) as well as IFN-β, IFN-γ, I-TAC, TRAIL, IL-17E/IL-25, IL-17F, IL-21, IL-22, IL-23, IL-27, IL-29/IFN-λ1, IL-31, and IL-33 (Biomarker Group 1), according to the manufacturer’s instructions. Plates were read with the MESO QuickPlex SQ 120 (Meso Scale Discovery), and analyte concentrations were calculated with the Discovery Workbench 4.0 (Meso Scale Discovery). The sensitivities of the assays were as follows: 0.21–0.62 pg/ml (IFN-γ), 0.01–0.17 pg/ml (IL-1β), 0.01–0.29 pg/ml (IL-2), 0.01–0.03 pg/ml (IL-4), 0.05–0.09 pg/ml (IL-6), 0.03–0.14 pg/ml (IL-8), 0.02–0.08 pg/ml (IL-10), 0.02–0.89 pg/ml (IL-12p70), 0.03–0.73 pg/ml (IL-13), 0.01–0.13 pg/ml (TNF), 3.1 pg/ml (IFN-β), 1.7 pg/ml (IFN-γ), 1.5 pg/ml (I-TAC), 0.66 pg/ml (TRAIL), 0.58 pg/ml (IL-17E/IL-25), 155 pg/ml (IL-17F), 1.2 pg/ml (IL-21), 0.13 pg/ml (IL-22), 1.4 pg/ml (IL-23), 9.6 pg/ml (IL-27), 1.2 pg/ml (IL-29/IFN-λ1), 7.3 pg/ml (IL-31), and 0.59 pg/ml (IL-33).

Cell culture supernatants and supernatants from TruCulture tubes were collected and stored as described above. The IMUBIND Tissue Factor ELISA kit (BioMedica Diagnostics, Windsor, Nova Scotia, Canada) was used to measure the concentrations of tissue factor (TF) in the supernatants according to the manufacturer’s instructions. Plates were read with the SpectraMax iD5 (Molecular Devices, San Jose, CA) and analyte concentrations were calculated with the SoftMax Pro 7 software (Molecular Devices). The sensitivity of the assay was 22.411 pg/ml.

Statistical analyses were performed by using the R language and environment. To compare flow cytometry data and cytokine intensities between study groups, linear mixed-effects models were fit to account for repeated observations from each patient. For cytokine mean fluorescence intensities, an offset was added to the data to ensure positive values, and the data were log2 transformed to improve normality. The flow cytometry data were modeled as proportions. A false discovery rate–adjusted p value <0.05 was considered statistically significant. For heat map representation of immunophenotyping results, flow cytometry data were transformed into z scores by subtracting the mean and dividing by the SD.

This study included peripheral blood samples collected from a largely African-American cohort of women in the third trimester of pregnancy as well as from healthy nonpregnant women of reproductive age. We exposed peripheral whole blood or PBMCs to inactivated SARS-CoV-2 particles, proteins, and peptide pools in vitro to thoroughly explore the systemic cellular immune responses triggered by this coronavirus in pregnant and nonpregnant women. Characterization of monocyte, T cell, and B cell subsets using flow cytometry revealed that both nonpregnant and pregnant women displayed strong responses to SARS-CoV-2 stimulation as indicated by altered phenotypes (presented in the Supplemental Extended Data). In this study, we solely focused on differences in cellular phenotypes between the nonpregnant and pregnant groups (shown as black asterisks throughout the figures) to investigate pregnancy-specific immune responses to in vitro challenge with SARS-CoV-2 particles and proteins/peptides.

Innate immune cells (e.g., neutrophils and monocytes) have been shown to undergo heightened activation during pregnancy (2830); therefore, it is likely that such immune cells act as a first line of defense against microbial infections during this period. We collected peripheral whole blood from pregnant and nonpregnant women and performed in vitro stimulation with inactivated SARS-CoV-2 particles to evaluate ROS production by neutrophils and monocytes (Fig. 1A). Consistent with prior reports (28, 29), pregnancy itself resulted in increased ROS production by peripheral monocytes and neutrophils (Fig. 1B). However, stimulation with SARS-CoV-2 particles did not modify ROS production by either pregnant or nonpregnant neutrophils and monocytes (Fig. 1B). These data show that the pregnancy-specific production of ROS by neutrophils and monocytes is not modified by SARS-CoV-2 particles.

FIGURE 1.

Monocyte and neutrophil responses to SARS-CoV-2 particles in pregnant and nonpregnant women. (A) Peripheral blood samples were collected from nonpregnant (n = 20, indicated in blue) and pregnant (n = 20, indicated in red) women to isolate PBMCs for in vitro stimulation with SARS-CoV-2 particles. Flow cytometry was performed to determine the expression of activation markers by monocyte subsets and neutrophils. (B, left) Flow cytometry gating strategy to measure the production of ROS in neutrophils and monocytes stimulated with SARS-CoV-2 particles (filled histograms) or control medium (open histograms). (B, right) Quantification of ROS production by neutrophils or monocytes after stimulation with SARS-CoV-2 particles (circles) or control medium (triangles). (C) Flow cytometry gating strategy for immunophenotyping of monocyte subsets. Monocytes were initially gated as viable CD14+ cells, followed by gating for classical (CD14hiCD16), intermediate (CD14hiCD16+), nonclassical (CD14loCD16+), and CD14loCD16 monocytes. (D) Heat map representations showing the expression of activation markers by total, intermediate, and classical monocytes after stimulation with SARS-CoV-2 particles. (E) Mean fluorescence intensity (MFI) of CD16 expression by total, intermediate, and classical monocytes from pregnant (red symbols) and nonpregnant (blue symbols) women in response to SARS-CoV-2 particles (circles) or control medium (triangles). *p < 0.05, **p < 0.01, ***p < 0.001. (+) Stimulated; (−) control.

FIGURE 1.

Monocyte and neutrophil responses to SARS-CoV-2 particles in pregnant and nonpregnant women. (A) Peripheral blood samples were collected from nonpregnant (n = 20, indicated in blue) and pregnant (n = 20, indicated in red) women to isolate PBMCs for in vitro stimulation with SARS-CoV-2 particles. Flow cytometry was performed to determine the expression of activation markers by monocyte subsets and neutrophils. (B, left) Flow cytometry gating strategy to measure the production of ROS in neutrophils and monocytes stimulated with SARS-CoV-2 particles (filled histograms) or control medium (open histograms). (B, right) Quantification of ROS production by neutrophils or monocytes after stimulation with SARS-CoV-2 particles (circles) or control medium (triangles). (C) Flow cytometry gating strategy for immunophenotyping of monocyte subsets. Monocytes were initially gated as viable CD14+ cells, followed by gating for classical (CD14hiCD16), intermediate (CD14hiCD16+), nonclassical (CD14loCD16+), and CD14loCD16 monocytes. (D) Heat map representations showing the expression of activation markers by total, intermediate, and classical monocytes after stimulation with SARS-CoV-2 particles. (E) Mean fluorescence intensity (MFI) of CD16 expression by total, intermediate, and classical monocytes from pregnant (red symbols) and nonpregnant (blue symbols) women in response to SARS-CoV-2 particles (circles) or control medium (triangles). *p < 0.05, **p < 0.01, ***p < 0.001. (+) Stimulated; (−) control.

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Monocytes are conventionally divided into several major subsets: classical (CD14hiCD16), intermediate (CD14hiCD16+), and nonclassical (CD14loCD16+), each of which displays distinct functionality (43). Thus, we next investigated whether SARS-CoV-2 particles differentially affected specific monocyte subsets in pregnant women. Few nonclassical monocytes were observed in our study samples; therefore, we focused on the phenotypes of the total, intermediate, and classical monocyte subsets (Fig. 1C). We evaluated the expression of specific surface markers, cytokines/chemokines, and receptors by total, intermediate, and classical monocytes in response to stimulation with SARS-CoV-2 particles (Fig. 1D, Supplemental Fig. 1A). Although SARS-CoV-2 particles reduced CD16 expression by total and intermediate monocytes from nonpregnant women, it tended to increase the expression of this marker by pregnancy-derived monocytes (Fig. 1E). CD16 expression in response to SARS-CoV-2 particle stimulation was elevated in total and intermediate monocytes from pregnant women compared to those from nonpregnant women (Fig. 1E). In addition, SARS-CoV-2 particle stimulation increased the typically minimal expression of CD16 by classical monocytes from both nonpregnant and pregnant women, suggesting that this coronavirus promotes a transition from classical to intermediate monocytes (Fig. 1E).

The SARS-CoV-2 virus has four major structural proteins: spike (S), membrane (M), envelope (E), and nucleocapsid (N) (44, 45) (Fig. 2A). The S protein (which includes an S1 and an S2 subunit) is critical for viral entry into host cells, thus it remains an attractive target for antiviral agents (45). The N protein is highly conserved, produced in abundance during infection, and strongly immunogenic (46). Therefore, we used the S and N proteins, the S1 and S2 subunits, and a mutant S1 (S1D614G) with increased infectious capacity (47, 48) to elucidate whether monocytes exhibit distinct responses to different SARS-CoV-2 proteins (Fig. 2A). Heat maps display the expression of surface markers, cytokines/chemokines, and receptors by stimulated total monocytes (Fig. 2B). Decreased expression of CD181 was observed on N protein–stimulated total monocytes from pregnant women compared to those from nonpregnant controls (Fig. 2C), and an increase in expression of CX3CR1 was observed in response to S protein stimulation (Fig. 2D). Given that intermediate monocytes appeared to be most activated by SARS-CoV-2 stimulation (Fig. 1E), we also determined the specific effects of viral proteins on this monocyte subset, as shown in the heat map representations (Fig. 3A). Consistent with responses observed in total monocytes (Fig. 2B, 2C), intermediate monocytes displayed decreased expression of CD181 in response to N protein (Fig. 3B) and increased CX3CR1 expression in response to S protein (Fig. 3C) in pregnant compared to nonpregnant women. Similarly, classical monocytes (Fig. 4A) derived from pregnant women also showed reduced expression of CD181 in response to N protein (Fig. 4B) but did not show changes in the expression of CX3CR1 in response to the S protein (Fig. 4C).

FIGURE 2.

Total monocyte response to SARS-CoV-2 proteins in pregnant and nonpregnant women. (A, left) Schematic representation of SARS-CoV-2 structure showing the S, M, E, and N proteins as well as the S protein subunits S1 and S2. (A, right) Peripheral blood samples were collected from nonpregnant (n = 20, indicated in blue) and pregnant (n = 20, indicated in red) women to isolate PBMCs for in vitro stimulation with SARS-CoV-2 proteins. Flow cytometry was performed to determine the expression of activation markers by total monocytes. (B) Heat map representations showing expression of activation markers by total monocytes after stimulation with SARS-CoV-2 proteins or the mutant S1 variant S1D614G. Mean fluorescence intensity (MFI) of CD181 expression (C) and CX3CR1 expression (D) by total monocytes from pregnant (red symbols) and nonpregnant (blue symbols) women in response to SARS-CoV-2 proteins (circles) or control (triangles). *p < 0.05, **p < 0.01. (+) Stimulated; (−) control.

FIGURE 2.

Total monocyte response to SARS-CoV-2 proteins in pregnant and nonpregnant women. (A, left) Schematic representation of SARS-CoV-2 structure showing the S, M, E, and N proteins as well as the S protein subunits S1 and S2. (A, right) Peripheral blood samples were collected from nonpregnant (n = 20, indicated in blue) and pregnant (n = 20, indicated in red) women to isolate PBMCs for in vitro stimulation with SARS-CoV-2 proteins. Flow cytometry was performed to determine the expression of activation markers by total monocytes. (B) Heat map representations showing expression of activation markers by total monocytes after stimulation with SARS-CoV-2 proteins or the mutant S1 variant S1D614G. Mean fluorescence intensity (MFI) of CD181 expression (C) and CX3CR1 expression (D) by total monocytes from pregnant (red symbols) and nonpregnant (blue symbols) women in response to SARS-CoV-2 proteins (circles) or control (triangles). *p < 0.05, **p < 0.01. (+) Stimulated; (−) control.

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FIGURE 3.

Intermediate monocyte response to SARS-CoV-2 proteins in pregnant and nonpregnant women. Peripheral blood samples were collected from nonpregnant (n = 20, indicated in blue) and pregnant (n = 20, indicated in red) women to isolate PBMCs for in vitro stimulation with SARS-CoV-2 proteins. Flow cytometry was performed to determine the expression of activation markers by intermediate monocytes. (A) Heat map representations showing the expression of activation markers by intermediate monocytes after stimulation with SARS-CoV-2 proteins or the mutant S1 variant S1D614G. Mean fluorescence intensity (MFI) of CD181 expression (B) and CX3CR1 expression (C) by intermediate monocytes from pregnant (red symbols) and nonpregnant (blue symbols) women in response to SARS-CoV-2 proteins (circles) or control (triangles). *p < 0.05. (+) Stimulated; (−) control.

FIGURE 3.

Intermediate monocyte response to SARS-CoV-2 proteins in pregnant and nonpregnant women. Peripheral blood samples were collected from nonpregnant (n = 20, indicated in blue) and pregnant (n = 20, indicated in red) women to isolate PBMCs for in vitro stimulation with SARS-CoV-2 proteins. Flow cytometry was performed to determine the expression of activation markers by intermediate monocytes. (A) Heat map representations showing the expression of activation markers by intermediate monocytes after stimulation with SARS-CoV-2 proteins or the mutant S1 variant S1D614G. Mean fluorescence intensity (MFI) of CD181 expression (B) and CX3CR1 expression (C) by intermediate monocytes from pregnant (red symbols) and nonpregnant (blue symbols) women in response to SARS-CoV-2 proteins (circles) or control (triangles). *p < 0.05. (+) Stimulated; (−) control.

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FIGURE 4.

Classical monocyte response to SARS-CoV-2 proteins in pregnant and nonpregnant women. Peripheral blood samples were collected from nonpregnant (n = 20, indicated in blue) and pregnant (n = 20, indicated in red) women to isolate PBMCs for in vitro stimulation with SARS-CoV-2 proteins. Flow cytometry was performed to determine the expression of activation markers by classical monocytes. (A) Heat map representations showing the expression of activation markers by classical monocytes after stimulation with SARS-CoV-2 proteins or the mutant S1 variant S1D614G. Mean fluorescence intensity (MFI) of CD181 expression (B) and CX3CR1 expression (C) by classical monocytes from pregnant (red symbols) and nonpregnant (blue symbols) women in response to SARS-CoV-2 proteins (circles) or control (triangles). *p < 0.05, **p < 0.01, ***p < 0.001. (+) Stimulated; (−) Control.

FIGURE 4.

Classical monocyte response to SARS-CoV-2 proteins in pregnant and nonpregnant women. Peripheral blood samples were collected from nonpregnant (n = 20, indicated in blue) and pregnant (n = 20, indicated in red) women to isolate PBMCs for in vitro stimulation with SARS-CoV-2 proteins. Flow cytometry was performed to determine the expression of activation markers by classical monocytes. (A) Heat map representations showing the expression of activation markers by classical monocytes after stimulation with SARS-CoV-2 proteins or the mutant S1 variant S1D614G. Mean fluorescence intensity (MFI) of CD181 expression (B) and CX3CR1 expression (C) by classical monocytes from pregnant (red symbols) and nonpregnant (blue symbols) women in response to SARS-CoV-2 proteins (circles) or control (triangles). *p < 0.05, **p < 0.01, ***p < 0.001. (+) Stimulated; (−) Control.

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To further refine our results, we performed stimulation of PBMCs by using pools of peptides with coverage of the complete M protein, the complete N protein, the S1 domain of the S protein, and the functional domains of the S protein, respectively (Supplemental Fig. 2A). Stimulation with these peptide pools was capable of inducing an immune response in both pregnant and nonpregnant women (Supplemental Extended Data). Yet, the heat maps displayed in Supplemental Fig. 2B illustrate the minimal difference in monocyte phenotypes and cytokine production between pregnant and nonpregnant women in response to each of the peptide pools.

Taken together, the above results indicate that SARS-CoV-2 modulates monocyte activation in a specific manner during pregnancy. Furthermore, such pregnancy-specific monocyte responses differ between exposure to viral particles and full-length S and N proteins, indicating a tailored monocyte response during SARS-CoV-2 challenge.

T cells, the primary cellular component of the adaptive immune system, are essential mediators of host antiviral responses (49). During pregnancy, the regulatory component of T cell immunity is enhanced (3336, 5052), whereas conventional Th functions are more tightly controlled (18, 5356); thus, we reasoned that T cell responses to SARS-CoV-2 Ags may differ between pregnant and nonpregnant women. PBMCs collected from nonpregnant and pregnant women were stimulated with SARS-CoV-2 particles as well as with the S and N proteins, the S1 and S2 subunits, S1D614G, and peptide pools (Fig. 5A), and the expression of multiple phenotypic, activation-associated, and functional markers by CD4+ and CD8+ T cells was determined (Fig. 5B, Supplemental Fig. 1B). T cells from nonpregnant and pregnant women showed minimal responsiveness toward SARS-CoV-2 proteins (Fig. 5C; Supplemental Extended Data). Apregnancy-specific increase in the expression of CD40L, which is essential for mediating macrophage and B cell activation (57), was observed on CD4+ T cells in response to the S2 subunit, given that treatment with the S1+S2 protein induced such a change, but not the S1 subunit alone (Fig. 5D). CD4+ T cells also exhibited responsiveness to inert SARS-CoV-2 particles as well as peptide pools, yet pregnancy-specific differences were not observed (Supplemental Fig. 3A, 3C; Supplemental Extended Data). Similarly, CD8+ T cells also displayed a pregnancy-specific increase in CD40L expression in response to stimulation with the S2 subunit (Fig. 6A, 6B), whereas no other distinct pregnancy-related changes were observed in response to SARS-CoV-2 particles or peptide pools (Supplemental Fig. 3B, 3D; Supplemental Extended Data). Together, these findings show that, whereas T cell activation is observed in response to SARS-CoV-2 Ags in pregnant and nonpregnant women, pregnancy itself only drives subtle differences in T cell activation.

FIGURE 5.

CD4+ T cell responses to SARS-CoV-2 Ags in pregnant and nonpregnant women. (A) Peripheral blood samples were collected from nonpregnant (n = 20, indicated in blue) and pregnant (n = 20, indicated in red) women to isolate PBMCs for in vitro stimulation with SARS-CoV-2 particles, proteins, or peptides. Flow cytometry was performed to determine the expression of activation markers by CD4+ and CD8+ T cells. (B) Representative gating strategy to identify the expression of activation markers on CD4+ and CD8+ T cells. (C) Heat map representations showing the expression of activation markers by CD4+ T cells after stimulation with SARS-CoV-2 proteins or the mutant S1 variant S1D614G. (D) Mean fluorescence intensity (MFI) of CD40L expression by CD4+ T cells from pregnant (red symbols) and nonpregnant (blue symbols) women in response to SARS-CoV-2 proteins (circles) or control (triangles). *p < 0.05. (+) Stimulated; (−) control.

FIGURE 5.

CD4+ T cell responses to SARS-CoV-2 Ags in pregnant and nonpregnant women. (A) Peripheral blood samples were collected from nonpregnant (n = 20, indicated in blue) and pregnant (n = 20, indicated in red) women to isolate PBMCs for in vitro stimulation with SARS-CoV-2 particles, proteins, or peptides. Flow cytometry was performed to determine the expression of activation markers by CD4+ and CD8+ T cells. (B) Representative gating strategy to identify the expression of activation markers on CD4+ and CD8+ T cells. (C) Heat map representations showing the expression of activation markers by CD4+ T cells after stimulation with SARS-CoV-2 proteins or the mutant S1 variant S1D614G. (D) Mean fluorescence intensity (MFI) of CD40L expression by CD4+ T cells from pregnant (red symbols) and nonpregnant (blue symbols) women in response to SARS-CoV-2 proteins (circles) or control (triangles). *p < 0.05. (+) Stimulated; (−) control.

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FIGURE 6.

CD8+ T cell responses to SARS-CoV-2 proteins in pregnant and nonpregnant women. (A) Heat map representations showing the expression of activation markers by CD8+ T cells after stimulation with SARS-CoV-2 proteins or the mutant S1 variant S1D614G. (B) Mean fluorescence intensity (MFI) of CD40L expression by CD8+ T cells from pregnant (red symbols) and nonpregnant (blue symbols) women in response to SARS-CoV-2 proteins (circles) or control (triangles). *p < 0.05. (+) Stimulated; (−) control.

FIGURE 6.

CD8+ T cell responses to SARS-CoV-2 proteins in pregnant and nonpregnant women. (A) Heat map representations showing the expression of activation markers by CD8+ T cells after stimulation with SARS-CoV-2 proteins or the mutant S1 variant S1D614G. (B) Mean fluorescence intensity (MFI) of CD40L expression by CD8+ T cells from pregnant (red symbols) and nonpregnant (blue symbols) women in response to SARS-CoV-2 proteins (circles) or control (triangles). *p < 0.05. (+) Stimulated; (−) control.

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The other major component of adaptive immunity is B cells, for which activation and class-switching are essential for the production of anti–SARS-CoV-2 IgM and IgG (58, 59). Therefore, we determined whether pregnancy-specific differences exist in B cell phenotypes in response to SARS-CoV-2 particles (Fig. 7A). We measured the expression of multiple phenotypic and activation-associated markers on peripheral B cells (Fig. 7B, Supplemental Fig. 1C) and noted a pregnancy-specific increase in the expression of CD22 coupled with decreased expression of CD24 in response to SARS-CoV-2 particle stimulation (Fig. 7C). We then examined whether changes in B cell surface marker expression were translated to a shift in the proportions of circulating B cell subsets. We examined the proportions of general B cell populations as well as multiple subsets of these cells (Fig. 7D). Notably, pregnancy was associated with an increase in the proportions of peripheral CD23+CD19+CD20+ B cells (Fig. 7E) and IgMIgDCD19+CD20+CD27+ B cells (Fig. 7F) compared to nonpregnant women. Only pregnancy-derived IgG+CD19+CD20+CD27+ B cells were expanded upon SARS-CoV-2 particle stimulation, thus the proportion of these cells was greater in pregnancy compared to the stimulated nonpregnant state (Fig. 7G). Peripheral CD23CD19+CD20+ B cells from both nonpregnant and pregnant women were decreased in response to stimulation, and such a decrease was more pronounced in pregnant women (Fig. 7H). By contrast, the CD24+CD38+CD19+CD20+ (Fig. 7I) and CD86+CD80+CD19+ CD20+ B cell (Fig. 7J) subsets were increased upon stimulation in both study groups, but this response was diminished in pregnant women compared to nonpregnant women. Interestingly, a stimulation-induced increase in the proportions of CD27+ CD43CD19+CD20+ (Fig. 7K) and CD27+IgD+CD19+CD20+ B cells (Fig. 7L) was observed only in nonpregnant individuals. Together, these data indicate that pregnancy is associated with altered proportions and phenotypes of peripheral B cells following challenge with SARS-CoV-2 particles and proteins/peptide pools.

FIGURE 7.

B cell responses to SARS-CoV-2 particles in pregnant and nonpregnant women. (A) Peripheral blood samples were collected from nonpregnant (n = 20, indicated in blue) and pregnant (n = 20, indicated in red) women to isolate PBMCs for in vitro stimulation with SARS-CoV-2 particles. Flow cytometry was performed to identify evaluate surface marker expression by B cell subsets. (B) Heat map representation showing the expression of surface markers by B cells after stimulation with SARS-CoV-2 particles. (C) Mean fluorescence intensity (MFI) of CD22 (left) and CD24 (right) expression by B cells from pregnant (red symbols) and nonpregnant (blue symbols) women in response to SARS-CoV-2 particles (circles) or control medium (triangles). (D) Heat map representation showing the proportions of B cell subsets within the parent population after stimulation with SARS-CoV-2 particles. Proportions of CD23+ (E), CD27+IgMIgD (F), CD27+IgG+ (G), CD23 (H), CD24+CD38+ (I), CD86+CD80+ (J), CD27+CD43 (K), and CD27+IgD+ (L) B cells from pregnant (red symbols) and nonpregnant (blue symbols) women in response to SARS-CoV-2 particles (circles) or control medium (triangles). *p < 0.05, **p < 0.01, ***p < 0.001. (+) Stimulated; (−) control.

FIGURE 7.

B cell responses to SARS-CoV-2 particles in pregnant and nonpregnant women. (A) Peripheral blood samples were collected from nonpregnant (n = 20, indicated in blue) and pregnant (n = 20, indicated in red) women to isolate PBMCs for in vitro stimulation with SARS-CoV-2 particles. Flow cytometry was performed to identify evaluate surface marker expression by B cell subsets. (B) Heat map representation showing the expression of surface markers by B cells after stimulation with SARS-CoV-2 particles. (C) Mean fluorescence intensity (MFI) of CD22 (left) and CD24 (right) expression by B cells from pregnant (red symbols) and nonpregnant (blue symbols) women in response to SARS-CoV-2 particles (circles) or control medium (triangles). (D) Heat map representation showing the proportions of B cell subsets within the parent population after stimulation with SARS-CoV-2 particles. Proportions of CD23+ (E), CD27+IgMIgD (F), CD27+IgG+ (G), CD23 (H), CD24+CD38+ (I), CD86+CD80+ (J), CD27+CD43 (K), and CD27+IgD+ (L) B cells from pregnant (red symbols) and nonpregnant (blue symbols) women in response to SARS-CoV-2 particles (circles) or control medium (triangles). *p < 0.05, **p < 0.01, ***p < 0.001. (+) Stimulated; (−) control.

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Finally, we investigated differences in mediator release by PBMCs from nonpregnant and pregnant women in response to SARS-CoV-2 particles (Fig. 8A). Baseline production of the proapoptotic TRAIL by pregnancy-derived PBMCs was diminished compared to those from nonpregnant women (Fig. 8B). SARS-CoV-2 stimulation enhanced the release of multiple cytokines, such as IFN-β, IFN-γ, TNF, IL-1β, IL-6, and IL-8, by PBMCs from nonpregnant and pregnant women (Fig. 8B). However, upon stimulation, PBMCs from pregnant women showed diminished release of IFN-β and IL-8 compared to those from nonpregnant individuals (Fig. 8B), suggesting restrained specific cytokine response to SARS-CoV-2 exposure in pregnant women. Several measured mediators displayed no changes with stimulation in nonpregnant or pregnant women (Supplemental Fig. 4A), including TF, a major component of the coagulation cascade that has been implicated in COVID-19 pathogenesis (60).

FIGURE 8.

Cytokine production by PBMCs exposed to SARS-CoV-2 particles or whole-blood TruCulture system containing S protein in pregnant and nonpregnant women. (A) Peripheral blood samples were collected from nonpregnant (n = 20, indicated in blue) and pregnant (n = 20, indicated in red) women to isolate PBMCs for in vitro stimulation with SARS-CoV-2 particles. Cytokine concentrations were then determined in culture supernatants. (B) Log10-transformed concentrations of IFN-β, IFN-γ, IL-1β, TNF, TRAIL, I-TAC, IL-2, IL-4, IL-6, IL-8, IL-10, IL-12p70, IL-13, IL-22, IL-23, IL-27, IL-29/IFN-λ1, and IL-33 in culture supernatants of PBMCs from nonpregnant (blue symbols) and pregnant (red symbols) women in response to SARS-CoV-2 particles (circles) or control medium (triangles). (C) Peripheral blood collected from nonpregnant (n = 14) and pregnant (n = 12) women was added to a whole-blood culture system (TruCulture) containing SARS-CoV-2 S protein. After culture, supernatants were collected for cytokine determination. (D) Log10-transformed concentrations of IFN-β, IFN-γ, IL-1β, TNF, TRAIL, IL-2, IL-4, IL-6, IL-8, IL-10, IL-17F, IL-27, IL-29/IFN-λ1, and IL-33 in whole-blood culture supernatants from pregnant (red symbols) and nonpregnant (blue symbols) women in response to S protein (circles) or control (triangles). *p < 0.05, **p < 0.01, ***p < 0.001. (+) Stimulated; (−) control.

FIGURE 8.

Cytokine production by PBMCs exposed to SARS-CoV-2 particles or whole-blood TruCulture system containing S protein in pregnant and nonpregnant women. (A) Peripheral blood samples were collected from nonpregnant (n = 20, indicated in blue) and pregnant (n = 20, indicated in red) women to isolate PBMCs for in vitro stimulation with SARS-CoV-2 particles. Cytokine concentrations were then determined in culture supernatants. (B) Log10-transformed concentrations of IFN-β, IFN-γ, IL-1β, TNF, TRAIL, I-TAC, IL-2, IL-4, IL-6, IL-8, IL-10, IL-12p70, IL-13, IL-22, IL-23, IL-27, IL-29/IFN-λ1, and IL-33 in culture supernatants of PBMCs from nonpregnant (blue symbols) and pregnant (red symbols) women in response to SARS-CoV-2 particles (circles) or control medium (triangles). (C) Peripheral blood collected from nonpregnant (n = 14) and pregnant (n = 12) women was added to a whole-blood culture system (TruCulture) containing SARS-CoV-2 S protein. After culture, supernatants were collected for cytokine determination. (D) Log10-transformed concentrations of IFN-β, IFN-γ, IL-1β, TNF, TRAIL, IL-2, IL-4, IL-6, IL-8, IL-10, IL-17F, IL-27, IL-29/IFN-λ1, and IL-33 in whole-blood culture supernatants from pregnant (red symbols) and nonpregnant (blue symbols) women in response to S protein (circles) or control (triangles). *p < 0.05, **p < 0.01, ***p < 0.001. (+) Stimulated; (−) control.

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We also evaluated mediator release in response to the SARS-CoV-2 S protein utilizing a whole-blood TruCulture system (42, 61) (Fig. 8C). The isolation of PBMCs is a useful technique for assaying the function of included immune cells, such as monocytes/macrophages and lymphocytes. However, the isolation of PBMCs excludes other immune cells, such as neutrophils, critical to the host response and obscures the effects of intercellular signaling in the peripheral blood milieu. Therefore, assessing both PBMC and whole-blood cultures can yield comprehensive insights into which cellular components are critical to the observed responses. Baseline whole-blood levels of IFN-γ and TRAIL were reduced in pregnant women compared to nonpregnant (Fig. 8D), whereas the concentrations of IL-1β, IL-8, IL-27, and IL-29/IFN-λ1 were increased. Upon stimulation, a pregnancy-specific elevation of IL-1β and IL-27 concentrations was observed together with reduced TRAIL (Fig. 8D). Interestingly, an increased whole-blood IFN-γ and IL-2 response to S protein was observed only in pregnant women (Fig. 8D). Several mediators, including TF, showed no response to stimulation in nonpregnant or pregnant women (Supplemental Fig. 4B).

Collectively, these results suggest that the basal levels of specific cytokines (e.g., IFN-γ, IL-1β, IL-18, TRAIL, and IL-27) in the maternal circulation are altered by pregnancy status; however, the S protein alone does not induce a unique immune response. Yet, the release of IFN-β and IL-8 by maternal mononuclear cells was diminished upon SARS-CoV-2 particle challenge, indicating that pregnancy imprints a differential cytokine response during coronavirus exposure.

Pregnancy itself is known to imprint specific immunological changes that can alter host responses to infection (37, 62), which is due, in part, to the dramatic physiologic changes and increasing energy demands that take place during this period (63). Mechanistic investigations of changes in cellular immune function during pregnancy have focused on differential responses to bacteria or their products, particularly endotoxin/LPS (37, 64, 65), given that bacterial infections are causally linked to pregnancy complications, such as preterm birth (6669). By contrast, mechanistic exploration of differential responses to viral exposure or infection during pregnancy to complement the vast body of clinical investigations is lacking. In this study, we evaluated the differential cellular responses to SARS-CoV-2 particle and protein/peptide challenge in PBMCs and whole blood isolated from nonpregnant and pregnant women. Recent reports on the SARS-CoV-2 pandemic indicated a pregnancy-specific neutrophilia among both infected and noninfected women (70), as observed in earlier studies unrelated to this virus (71). Moreover, neutrophils display a hyperactivated state during pregnancy (72), which could contribute to their elevated ROS production and potentially explain why stimulation with SARS-CoV-2 particles did not alter such function in the current study. Indeed, the elevated activation status/ROS production of neutrophils could result in increased COVID-19 severity in pregnant women (73), given that in silico analysis of peripheral neutrophils from nonpregnant cases of severe disease revealed upregulation of transcripts associated with oxidative stress (74). However, whether pregnancy-driven neutrophil activation is implicated in the severity of SARS-CoV-2 infection remains unclear.

Monocytes are primary participants in the host defense against viral infection (75). Under steady-state conditions, circulating classical, intermediate, and nonclassical monocytes form a continuum of sequential transition, and interruption of this cycle during a systemic inflammatory response is counteracted by the early release of classical monocytes from the bone marrow (76). Consistently, significant monocyte expansion has been reported in cases of severe COVID-19 among patients who were admitted to the intensive care unit (77, 78). The gene expression profiles of monocytes from cases of severe COVID-19 display highly dysregulated gene expression profiles (77, 79), which are indicative of enhanced cell activation and a migratory phenotype (79). Such intense alterations in monocyte phenotypes observed in COVID-19 patients are suggestive of emergency myelopoiesis (80) [i.e., the emergency activation of bone marrow hematopoiesis to accommodate demand for myeloid cells (81)]. Notably, the predominant cellular infiltrate in bronchoalveolar fluid obtained from COVID-19 patients is monocytes, and the abundance of these cells has been correlated with disease severity (82). In line with the abovementioned evidence, our current data support the accelerated transition and activation of monocytes in response to SARS-CoV-2 particles. Notably, monocytes derived from pregnant women showed elevated expression of CD16 in response to stimulation compared to those derived from nonpregnant women, suggesting enhanced activation and/or a more intensive transition process. Although monocytes from both pregnant and nonpregnant women displayed increased expression of activation markers in response to SARS-CoV-2 particles, the pregnancy-specific increase in CD16 expression could signify enhanced monocyte activation/transition that may contribute to the more severe disease progression observed in pregnant women (68, 83). Similarly, a prior study of pregnant women with mild COVID-19 noted a tendency for reduced classical monocytes and elevated intermediate monocytes compared to nonpregnant women (84), suggesting that a similar phenomenon is observed in women with severe COVID-19.

Interestingly, a previous investigation reported that the exposure of monocytes from healthy donors to SARS-CoV-2 N or S proteins resulted in increased expression of IL6, IL1B, and IL10 as well as enhanced IL-6 release, suggesting the potential contribution of these cells to the uncontrolled production of inflammatory mediators observed in severe COVID-19 cases (85). Similarly, in this study, we show that stimulation with SARS-CoV-2 proteins provokes an extensive in vitro inflammatory response in monocytes isolated from pregnant and nonpregnant women. Notably, the effects of N protein exposure on monocytes have been shown to be more intense than those of the S protein (85), potentially due to the highly immunogenic nature of such protein (46). This is consistent with our data showing that monocyte responses are greatest upon exposure to the N protein followed by the S protein, regardless of pregnancy status. Furthermore, we observed pregnancy-specific alteration of surface expression of monocyte receptors such as CD181 (the IL-8 receptor) and CX3CR1 upon stimulation with the N and S proteins, respectively, suggesting that specific viral proteins could modulate monocyte chemotactic responses in pregnant women exposed to SARS-CoV-2.

Monocyte responses in pregnant and nonpregnant women were also evaluated in response to peptide pools corresponding to the SARS-CoV-2 N, S, and M proteins. Monocytes derived from both study groups show extensive phenotypic changes in response to peptide exposure; yet, no intergroup differences in response were observed. A previous study, in which the authors investigated T cell responses to HLA-DR– and HLA-A*24–restricted peptides in convalescent COVID-19 patients, identified epitopes with promiscuous or specific reactivity, respectively, that could govern long-term immunity (86). Therefore, the detection of specific SARS-CoV-2 peptides by memory T cells could be an important component of the immune response against secondary infection. However, in the current study, peripheral immune cells were isolated from pregnant and nonpregnant women and exposed to SARS-CoV-2 peptides in an in vitro setting, which may not elicit comparable responses to those induced by full-length proteins or viral particles.

SARS-CoV-2 infection is linked to lymphopenia, which includes reduced systemic proportions of T cells (8789), and we recently reported a similar phenomenon in pregnant women infected with SARS-CoV-2 (90). Such a phenomenon is reflected in the peripheral blood transcriptome of COVID-19 patients, in which upregulated transcripts were enriched for innate immune process, whereas those downregulated included T cell terms (87). Longitudinal immune profiling of severe COVID-19 patients demonstrated excessive T cell activation, potentially indicating a compensatory increase in effector functionality to overcome diminished numbers (88, 89), which is consistent with the pregnancy-specific increase in T cell–associated cytokines such as IL-2 and IFN-γ observed in the current study. These observations raise the critical question of how changes in the peripheral T cell compartment induced by SARS-CoV-2 infection interact with those that occur naturally in pregnant women. During normal pregnancy, immunological adaptations include the specific modulation of T cell signatures in the maternal circulation that increase throughout the third trimester and in normal labor at term (72, 9193), which may reflect changes occurring in the T cell compartment in the placental tissues (94). Indeed, we recently showed that the single-cell transcriptomic profiles of T cells in the chorioamniotic membranes from women with COVID-19 during pregnancy partially overlap with those of peripheral T cells from hospitalized COVID-19 patients (90). Therefore, immunological modulation by SARS-CoV-2 exposure is not limited to the subtle changes in peripheral T cell phenotypes observed in this study, but also extends to the maternal–fetal interface.

Extensive research has shown that COVID-19–induced lymphopenia includes a significant decrease in multiple subsets of circulating B cells with the exception of plasmablasts, which are increased in infected patients with SARS-CoV-2 and correlated with disease severity (87, 9597). In the context of pregnancy, it has also been shown that the numbers and functions of B cells in the peripheral blood of pregnant women with COVID-19 differ from those of nonpregnant women with COVID-19 (98). In this study, we found that B cell expression of CD22, which is a negative regulator of BCR signaling (99), is increased in pregnant compared to nonpregnant women upon stimulation with SARS-CoV-2 particles. This is in line with a previous report showing the reduced expression of molecules associated with BCR signaling, such as RAC2, CD79B, PTPRC, and BLNK in pregnant women with COVID-19 compared to nonpregnant patients (98). We also observed a pregnancy-specific reduction in the unswitched memory-like and transitional-like B cell subsets, whereas switched memory-like B cells were more expanded in pregnant women upon stimulation. These results partially overlap with prior studies noting that the severity of COVID-19 was linked to reduced abundance of unswitched memory B cells (95) and may be indicative of an impaired B cell response in pregnant women with SARS-CoV-2 infection. Memory B cells may undergo accelerated class switching in response to SARS-CoV-2 in pregnant women, which could explain the diminished proportions of unswitched memory-like B cells and elevated proportions of switched memory-like B cells compared to nonpregnant women. A limitation of our B cell analyses is that we did not explore the Ig profiles of these cells in response to in vitro stimulation with SARS-CoV-2 particles or proteins. Yet, multiple investigations focused on in vivo B cell responses in pregnant women infected with SARS-CoV-2 (39, 100, 101), therefore diminishing the need to evaluate this concept by using viral particle challenge.

It is well documented that severe COVID-19 is characterized by the onset of a massive cytokine storm (102104). Indeed, the systemic concentrations of proinflammatory cytokines have been used in the clinical setting to predict disease severity (105), and multiple proposed treatments for COVID-19 were directed toward mechanisms of cytokine release (104, 106). In this study, we found that stimulation with SARS-CoV-2 particles increased the release of multiple cytokines by peripheral leukocytes from both pregnant and nonpregnant women, with the production of IFN-β and IL-8 diminishing in pregnancy. IFNs are a critical component of antiviral defense (107, 108). Type I IFNs, including IFN-β, bind to their receptors (IFNAR) and induce the transcription of IFN-stimulated genes, which restrict viral replication through different mechanisms (109, 110). A recent study reported no differences in the plasma concentrations of IFN-β between pregnant women who had recovered from SARS-CoV-2 infection and those with ongoing or recent infection (111); however, a direct comparison of systemic IFN-β levels between pregnant and nonpregnant women has not been made. Previous research demonstrated the importance of the timing of the IFN response in relation to viral replication, as a central determinant of the observed outcomes. Although the early induction of type I IFN confers maximum protection, a delayed response will not only fail to control viral replication but can also lead to inflammation and tissue damage (110). Therefore, the current finding that IFN-β release in response to SARS-CoV-2 is diminished during pregnancy could indicate a delayed IFN response and thus contribute to a more severe phenotype in pregnant women.

IL-8, also known as CXCL8, is a proinflammatory chemokine associated with neutrophil chemotaxis as well as with the activation of such innate immune cells (112). In the clinical setting, IL-8 has been demonstrated to play a role in the pathophysiology of obstetrical syndromes as evidenced by altered concentrations of this chemokine in the peripheral blood of women with preeclampsia (113, 114) as well as in the amniotic fluid of women who underwent preterm labor and birth (115, 116), where it is highly correlated with the presence of acute histologic chorioamnionitis (116). Indeed, IL-8 in cervical fluid has been used as a marker of intra-amniotic inflammation in a noninvasive point-of-care test to monitor women with preterm prelabor rupture of the membranes (117). In addition to perinatal complications, elevated amniotic fluid IL-8 concentrations are also associated with adverse neonatal outcomes, such as bronchopulmonary dysplasia (118). Thus, modulation of systemic IL-8 concentrations by SARS-CoV-2 could be relevant for the care of infected pregnant women. In the current COVID-19 pandemic, IL-8 was shown to be increased in the peripheral blood from individuals with mild-moderate COVID-19 compared to healthy controls as well as in patients with severe COVID-19 compared to those presenting mild-moderate disease (119). A prior report did not find differences in systemic IL-8 concentrations between pregnant women who had recovered from SARS-CoV-2 infection and currently infected or convalescent pregnant women (111). Similarly, no differences were reported between pregnant and nonpregnant women with COVID-19 (70); however, only four patients were included in the latter study group. By contrast with the above studies, in this study, we show differential release of IL-8 from peripheral leukocytes of pregnant and nonpregnant women in response to SARS-CoV-2 viral particle exposure. Such differences could be due, in part, to the altered neutrophil biology taking place during pregnancy, which is exacerbated by systemic inflammation (29). As pregnant women with COVID-19 are at increased risk of developing preeclampsia (7, 9), and as the pathophysiology of preeclampsia involves endothelial dysfunction related with changes in IL-8 (113, 114), it is tempting to speculate that the altered SARS-CoV-2 particle-induced release of IL-8 from the PBMCs of pregnant women may contribute to the intravascular immune response implicated in the pathogenesis of preeclampsia.

The findings described in this study must be interpreted with caution due to the in vitro nature of the study. Patients who self-reported as testing positive for SARS-CoV-2, having COVID-19, or receiving a vaccine were excluded from the study; yet, we were unable to rule out possible undiagnosed asymptomatic infection in these patients. Moreover, previous studies indicated that T cells from patients with prior coronavirus infection were cross-reactive with SARS-CoV-2 epitopes (120). In the current study, the prior history of each patient was not available, thus any potential contribution of T cell cross-reactivity to the results reported herein could not be accounted for. Lastly, it is worth mentioning that there are potential confounding factors between pregnant and nonpregnant women that may influence the reported results, such as pregnancy-related medication use, dietary self-supplementation, and lifestyle changes, among others. Therefore, further in vivo and ex vivo investigations of the effects of pregnancy on the maternal immune response to SARS-CoV-2 are needed.

In summary, the in vitro model used in this study allows for the evaluation of peripheral immune responses triggered by SARS-CoV-2 in pregnant and nonpregnant women. Overall, our current findings are consistent with those that were recently reported in our ex vivo investigation of pregnant women with COVID-19, the majority of which presented a mild form of the disease (90). However, our model does not account for critical changes taking place in maternal organs as well as those at the maternal–fetal interface, which have the largest impact on pregnancy outcome. Thus, different in vitro systems must be developed to fully simulate the immunological responses taking place in pregnant women with severe COVID-19, given that such patients are at greater risk of adverse perinatal consequences due to SARS-CoV-2 infection.

We thank Rona Wang from the Translational Science & Biomarkers Unit of the Perinatology Research Branch for carrying out some molecular assays and George Schwenkel and Megan Faucett from the Clinical Laboratory of the Perinatology Research Branch for coordinating sample collection. The authors also thank the physicians, nurses, and research assistants from the Center for Advanced Obstetrical Care and Research, Intrapartum Unit, and the Perinatology Research Branch Clinical Laboratory for help with collecting samples.

This work was supported by the Perinatology Research Branch, Division of Obstetrics and Maternal-Fetal Medicine, Division of Intramural Research, Eunice Kennedy Shriver National Institute of Child Health and Human Development, National Institutes of Health, U.S. Department of Health and Human Services under Contract HHSN275201300006C (to R.R.). This research was also supported by the Wayne State University Perinatal Initiative in Maternal, Perinatal and Child Health (to N.G.-L. and A.L.T.). R.R. has contributed to this work as part of his official duties as an employee of the United States Federal Government.

The online version of this article contains supplemental material.

Abbreviations used in this article:

     
  • COVID-19

    coronavirus disease 2019

  •  
  • E

    envelope

  •  
  • M

    membrane

  •  
  • N

    nucleocapsid

  •  
  • ROS

    reactive oxygen species

  •  
  • S

    spike

  •  
  • SARS-CoV-2

    severe acute respiratory syndrome coronavirus 2

  •  
  • TF

    tissue factor

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The authors have no financial conflicts of interest.

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