Infections during pregnancy can expose the fetus to microbial Ags, leading to inflammation that affects B cell development. Prenatal fetal immune priming may have an important role in infant acquisition of pathogen-specific immunity. We examined plasma proinflammatory biomarkers, the proportions of various B cell subsets, and fetal priming to tetanus vaccination in cord blood from human United States and Kenyan neonates. United States neonates had no identified prenatal infectious exposures, whereas Kenyan neonates examined had congenital CMV or mothers with prenatal HIV or Plasmodium falciparum or no identified infectious exposures. Kenyan neonates had higher levels of IP-10, TNF-α, CRP, sCD14, and BAFF than United States neonates. Among the Kenyan groups, neonates with prenatal infections/infectious exposures had higher levels of cord blood IFN-γ, IL-7, sTNFR1, and sTNFR2 compared with neonates with no infectious exposures. Kenyan neonates had greater proportions of activated memory B cells (MBC) compared with United States neonates. Among the Kenyan groups, HIV-exposed neonates had greater proportions of atypical MBC compared with the other groups. Although HIV-exposed neonates had altered MBC subset distributions, detection of tetanus-specific MBC from cord blood, indicative of fetal priming with tetanus vaccine given to pregnant women, was comparable in HIV-exposed and non–HIV-exposed neonates. These results indicate that the presence of infections during pregnancy induces fetal immune activation with inflammation and increased activated MBC frequencies in neonates. The immunologic significance and long-term health consequences of these differences warrant further investigation.

Infections such as HIV, CMV, and malaria are common during pregnancy in sub-Saharan Africa and are associated with maternal inflammation and immune activation. These infections can be associated with negative pregnancy and birth outcomes such as maternal and fetal anemia, preterm birth, and low birth weight. However, there is a spectrum of clinical manifestations with many neonates having no apparent clinical consequences. In the context of a successful term pregnancy, how these infections affect fetal B cell development or whether these lead to fetal immune activation is poorly understood.

The human fetus is generally thought to have a functionally immature immune system with increased susceptibility to infection (1, 2). However, research has shown that T and BCR repertoires are diverse by the end of the second trimester (3, 4). Numerous reports have demonstrated fetal immune priming to foreign Ags that cross the placenta and may modulate neonatal/infant immune responses. Neonatal T cell recall responses are elicited by HIV, CMV, and malaria Ags (511). Evidence for transplacental priming of fetal B cells has been shown in studies examining cord blood for Ag-specific IgM and IgE, which cannot cross the placenta from the maternal circulation and are therefore of fetal origin (1114). Several studies have shown that fetal immune priming might confer postnatal protection against infection (6, 15, 16), whereas others suggest that this may lead to the development of allergies (1719), increased risk of infections (10, 2022), and decreased protective immunity to vaccinations (23, 24). The biological processes behind the varied consequences of prenatal immune priming are yet to be fully understood.

B cells are multifunctional lymphocytes involved in development of acquired immunity to many pathogens. Apart from their role in humoral immune defense, B cells also act as potent APCs, produce numerous cytokines, and contribute to T cell regulation. Early B lymphopoiesis and peripheral B cell maturation is regulated by several transcriptional factors and cytokines that act at specific time points, such as IL-7, IP-10, and BAFF (25). B cells can be classified by surface immunophenotyping into distinct subsets according to their state of maturation and differentiation. CD27 expression characterizes memory B cells (MBC) (26). MBC are thought to be a heterogenous population with classic isotype-switched MBC (CD27+IgG+IgD) (the predominant responders to secondary Ag challenge [27]). When activated, these MBC are characterized as activated MBC (CD27+CD21). Within the last decade, a population of hyporesponsive MBC characterized by CD27CD21 called “exhausted” or “atypical” MBC were found to be expanded in individuals with infections such as HIV, malaria, and hepatitis C virus (2831). This population has evidence of somatic hypermutation consistent with classic MBC but with variable Ab production after stimulation (3235). The expansion of atypical MBC is typical of some infections associated with delayed development of immunologic memory. Finally, nonswitched MBC (IgD+CD27+) frequencies have been found to be lower in infants from malaria-endemic regions (36). This population includes IgM+IgD+CD27+ MBC similar to marginal zone MBC and has an important role in protection against encapsulated bacteria (27, 37).

The focus of our study is to understand how prenatal infections, such as HIV, CMV, and malaria, affect fetal B cell maturation, activation, and memory formation. We hypothesize that neonates born to mothers with prenatal infections (CMV) or infectious exposures (HIV or malaria) will have increased proinflammatory molecules associated with B cell activation and maturation in cord blood that result in more activated and responsive B cells. We tested this hypothesis by examining plasma cytokines in United States cord blood and Kenyan maternal–neonatal pairs and B cell subset frequencies in United States and Kenyan cord blood. Kenyan pregnant women enrolled included women who had HIV or malaria, or neonates with congenital CMV, or none of these three infections. Additionally, we examined fetal priming to tetanus vaccination in HIV-exposed and non–HIV-exposed neonates.

Healthy pregnant mothers were recruited from antenatal clinics at Msambweni District Hospital and Port Reitz Hospital in Coast Province, Kenya from 2005 to 2010. Per Kenya Ministry of Health national policy, women received sulfadoxine-pyrimethamine beginning in the second trimester (for a total of three doses at intervals of 1 mo or more) in addition to iron, folic acid, and bed nets as part of routine care. All participants were tested for HIV-1 and HIV-2 as part of the volunteer counseling and testing program and, if positive, were treated according to Kenyan Ministry of Health policy. At the first antenatal visit, consented women were tested for malaria, geohelminths, and urinary schistosomiasis and were treated if indicated. Schistosomiasis was not treated until after delivery according to the Kenyan guidelines at the time. Demographic information was obtained, and a physical examination was performed. Pregnant women with known medical disorders contributing to fetal growth restriction, placental dysfunction, twin pregnancy, and prematurity were excluded. Participating women who delivered a live term infant had maternal venous and cord blood obtained as previously described (20). Birth weight was obtained immediately after delivery and birth length and head circumference obtained within 24 h of delivery. United States cord blood was collected from healthy term neonates under a discarded tissue protocol. No further information regarding clinical status of the United States neonate or mother was collected. United States adult volunteers (equal number of men and women) also donated peripheral blood for these studies. The study was approved by the University Hospital of Cleveland Medical Center Institutional Review Board and the Kenyan Medical Research Institute/Ethical Review Committee.

Cord blood was obtained from the umbilical vein of the placenta immediately after delivery. Peripheral venous blood was obtained by venipuncture from Kenyan and United States adults. All samples were anticoagulated with heparin and processed within 4 h. Cord blood mononuclear cells (CBMC) were then isolated by Ficoll-Hypaque density gradient centrifugation as previously described (38). Plasma was obtained after centrifugation. The mononuclear cells were cryopreserved with 10% DMSO and heat-inactivated FBS at −80°C for 24 h, then transferred to liquid nitrogen (38). PBMC from United States healthy adult volunteers were processed in the same way.

Our goal was to interrogate ∼10 samples from each comparator group. We first selected sample sets that had paired maternal and cord blood plasma available. We then processed CBMC samples that, upon thawing, had ≥70% cellular viability. Because of a large number of samples having poor viability, fewer than 10 samples in some groups were available. All viable available samples were examined and data presented.

Kenyan HIV-exposed neonates.

HIV testing of pregnant women by ELISA was performed by the Kenyan Ministry of Health as was management of antiretroviral regimens and clinical response. CD4 counts were not available for all HIV+ women, and HIV viral loads were not measured. All HIV+ women received zidovudine, lamivudine, and nevirapine during pregnancy until delivery. Neonates of HIV+ mothers were term (>37 wk gestational age), and all were determined to be HIV− by PCR at 12 mo of age.

Kenyan malaria-exposed neonates.

Pregnant women were screened for Plasmodium falciparum infection at their first antenatal visit (peripheral blood) and at delivery (peripheral, placental intervillous, and cord blood). Malaria parasites were detected by 1) thick and thin blood smears stained with Geimsa and examined by light microscopy (20) and 2) DNA extracted from blood and subjected to PCR-ligase detection reaction/fluorescent microsphere assay as previously described (39). To determine if Kenyan neonates were primed to P. falciparum Ags in utero, CBMC were coincubated with P. falciparum merozoite surface protein 1 (MSP1) peptides and PfP0 (a merozoite surface protein [40, 41]) T cell epitope peptides and T cell recall responses measured by ELISpot and cytokine production in supernatants measured by ELISA for IFN-γ, IL-2, IL-5, and IL-13 as previously described (20). CBMC coincubated with PMA or media only were used as positive and negative controls. A newborn was considered to be sensitized to P. falciparum in utero when one of the following three conditions were met: 1) by IFN-γ ELISPOT, there were more than four cytokine-secreting cells per 106 CBMC in response to P. falciparum Ag peptide(s) and no secreting cells were detected in negative control wells; 2) by IFN-γ ELISPOT, in cases where cytokine-secreting cells were observed in negative control wells, the number of spots generated by P. falciparum peptide-driven CBMC was 2-fold greater than control wells; 3) by ELISA for IFN-γ, IL-2, IL-5, or IL-13, net cytokine production by CBMC in response to P. falciparum Ag peptides was at least 2-fold greater than that of negative control wells (20). Thus neonates classified as “malaria exposed” examined in this study were exposed to malaria in utero and had resultant fetal T cell immune priming to these Ags.

Kenyan neonates with congenital CMV.

Congenital CMV is typically diagnosed by culturing infant urine for CMV virus or PCR detection of CMV, which was not possible at the study sites. An alternative method of congenital CMV detection is the presence of CMV IgM Abs in cord blood. Maternal IgM does not cross the placenta (42). CMV IgM is typically used to diagnose primary CMV infection in, for example, pregnant women (43). CMV IgM detectable in cord blood has been used to diagnose congenital CMV but with lower sensitivity and specificity than culture of CMV from urine or PCR detection of CMV from cord blood (14, 44). We used a microsphere-based multiplex method to quantify CMV IgM and IgG Abs in cord blood plasma. Briefly, 8 μg of the CMV internal matrix protein pp65 (Miltenyi Biotech) was coupled to carboxylated microspheres (Luminex, Austin, TX) using the manufacturer’s protocol and as previously described (45, 46). Cord blood plasma at a 1:100 dilution was incubated with 500 CMV protein–conjugated microspheres per well. After washing, CMV-specific IgM was detected with a 1:50 dilution of goat F(ab′)2 anti-human IgM R-PE conjugate (Southern Biotech). CMV-specific IgG was detected with a 1:100 dilution of goat F(ab′)2 anti-human IgG R-PE conjugate (Jackson ImmunoResearch). At least 75 protein-conjugated microspheres per well were then acquired on a Bioplex reader (Bio-Rad, Hercules, CA) and results expressed as the mean fluorescence intensity (MFI). A CMV IgM of >3000 MFI was considered positive. Negative controls had background MFIs of <100 MFI. CMV IgG was also measured. For this study, four participants were included who were CMV IgM positive. Two of these four had CMV IgM levels that were higher than their CMV IgG levels. For the entire cohort tested with this method, 67 neonates had positive CMV IgM out of 521 tested for congenital CMV, giving a prevalence of 12.9% for this population, which is similar to published studies from Kenya (47, 48). Of the HIV-exposed cord blood samples used for the comparative infections flow cytometry/plasma cytokine study (n = 13), two were also CMV IgM+. We elected to keep them in the HIV group, as potentially more confounders (antiretroviral exposure, maternal CD4 count) affect this group than the CMV group. All CMV IgM+ neonates had no detected clinical features of congenital CMV. Neonatal hearing was not tested, as testing was not available. This particular method of detecting CMV IgM was not validated with CMV viral culture nor PCR, which is a limitation to this study.

Kenyan nonexposed neonates.

Mothers of these neonates were HIV negative, had no evidence of any malaria infections during pregnancy, and cord blood was CMV IgM negative. We use the term “nonexposed” to detected infections, realizing that other maternal infections such as geohelminth or bacterial urinary tract infections may have occurred.

United States nonexposed neonates.

No infections were known to be associated with these pregnancies. Plasma was negative for CMV IgM. Mothers were HIV−.

Data from Kenyan HIV-exposed and malaria-exposed neonates is combined with data from Kenyan neonates with congenital CMV and compared with neonates for whom none of these infections/infectious exposures were detected. This group is labeled “Kenyan all infections/infectious exposure group,” and in figures, a shortened label of “K all inf” is used to refer to this group.

Plasma cytokines (IFN-γ, IP-10, TNF-α, IL-4, IL-6, IL-7, IL-9, IL-10, IL-12 (p40), IL-12 (p70), IL-17α, sTNFR1, and sTNFR2) from cord blood and peripheral blood plasma were measured using a Millipore MILLIPLEX MAP Human Cytokine/Chemokine Magnetic Bead Panel per manufacturer’s instructions (EMD Millipore, Billerica, MA). Two CMV+ cord blood plasma samples were not available for this cytokine assay. Plasma BAFF and CRP concentrations from cord blood and peripheral blood were measured using the Quantikine ELISA Human BAFF Immunoassay and Quantikine ELISA Human CRP Immunoassay (R&D Systems, Minneapolis, MN) per manufacturer’s instructions.

Cryopreserved CBMC were thawed and rested. The viability of the cells was initially verified using trypan blue. Samples with poor viability (>30% dead cells) were not examined. Prior to surface staining, thawed cells were stained with LIVE⁄DEAD Fixable Red Dead Cell Stain (Invitrogen) for 30 min, then washed and surface stained on ice in a final 100 μl of staining volume using combinations of the following commercially conjugated mouse anti-human mAb for analysis of B or T cell phenotypes. Six-color panels were developed to differentiate B cell subpopulations; panel 1: CD19 PErCP-Cy5.5, CD27 PeCy-7, IgM FITC, IgD allophycocyanin-H7, CD10 PE, CD21 Pe-Cy7; panel 2: CD19 PErCP-Cy5.5, CD27 PeCy-7, TLR2 PE, CD5 allophycocyanin, IgM FITC, IgD allophycocyanin-H7. Panel 3 was used to identify CD4 and CD8 populations (CD45 allophycocyanin, CD3 Alexa Fluor 700, CD8 PerCP, CD4 FITC). Cells were washed with PBS and 2% FBS, fixed with 1% paraformaldehyde for 30 min, and analyzed with a BD LSRII (BD Biosciences, San Jose, CA) flow cytometer. Uncompensated data were collected for all experiments. Compensation corrections were computed, and analyses were carried out using FlowJo (Tree Star, Ashland, OR). For each panel, at least 100,000 events were collected and gated to identify lymphocytes (forward scatter [FSC] versus side scatter [SSC]) and exclude dead cells (Red ViViD). We defined immature B cells as CD19+CD10+CD21+IgD+, transitional B cells as CD19+CD10+CD21+IgD, mature naive B cells as CD19+CD27CD21+CD10, classic isotype-switched MBC as CD19+CD27+CD21+IgD, activated MBC as CD19+CD27+CD21IgD, and atypical MBC as CD19+CD27CD21CD10 as previously defined (30). We also evaluated the presence of TLR2 and CD5 in the B cell populations as part of stimulation assays. Fig. 1 shows an example of the gating strategy that was used to identify B cell subpopulations (e.g., classic isotype-switched MBC). The source of Abs is as follows: CD19 PErCP-Cy5.5, CD27 PeCy-7, CD4 FITC, TLR2 PE (eBiosciences, San Diego, CA); IgM FITC, CD8 PerCP, CD5 allophycocyanin (BioLegend, San Diego, CA); IgD allophycocyanin-H7, CD10 PE, CD21 Pe-Cy7, CD45 allophycocyanin, CD3 Alexa Fluor 700 (BD Biosciences).

Cryopreserved CBMC and PBMC were thawed and divided in half into wells containing media alone or media with a stimulation mixture consisting of PWM (50 ng/ml) (Sigma-Aldrich, St Louis, MO), SAC protein A (1:10,000) (Sigma-Aldrich), and CpG ODN 2006 (5 μg/ml) (Operon, Huntsville, AL). The cells, with and without stimulation, were cultured for 12 h at 37°C in 5% CO2. They were then washed thoroughly with PBS and 2% FBS, stained, and analyzed by flow cytometry as described above.

Protein biotinylation and formation of Ag–quantum dot (QD) complexes was performed using diphtheria toxin CRM197 (DT; List Biological Labs, CA), tetanus toxoid (TT; List Biological Labs), and human serum albumin (HSA; Sigma-Aldrich; used as a negative control) that were biotinylated using a ChromaLink Biotin Kit (Solulink, CA) according to the manufacturer’s instructions. Briefly, 100 μg (1 mg/ml) DT diluted in 10 mM sodium phosphate pH 7.5/5% α lactose monohydrate (Sigma-Aldrich) buffer was biotinylated with 7.5-fold excess of biotin reagent, 100 μg (1 mg/ml) of TT diluted in 1× PBS was biotinylated with a 7-fold excess of biotin reagent, and 100 μg (1 mg/ml) HSA in 1× PBS was biotinylated with a 5-fold excess of biotin, which accomplished the desired labeling of one biotin molecule to one primary amine per protein. Biotinylated TT and DT were validated by comparing biotinylated and nonbiotinylated Ag in parallel ELISA and 5-d B cell ELISpot assays using plasma and PBMC from vaccinated North American adults. No discernible differences were detected, underscoring that biotinylation does not affect B cell epitopes (data not shown).

All Ag-QD complexes were prepared fresh for each experiment. Biotinylated Ag was incubated with StrepAvidin-QD605, QD655, or QD705 (Invitrogen Life Technologies, NY) at a 25:1 (Ag:QD) molar ratio. For DT-QD complex formation, the QD was diluted in 10 mM sodium phosphate pH 7.5/5% α lactose monohydrate (Sigma-Aldrich). A 1/10 volume of this was added to biotinylated DT on ice every 10 min. For TT-QD and HSA-QD, the QD was diluted in PBS, and 1/10 volume of this was added to biotinylated TT or HSA on ice every 10 min. Before cell staining, Ag-QD complex preparations were centrifuged for 10 min at maximum speed to remove aggregates. For TT-MBC frequency determination in CBMC, the TT C-fragment (TTc) with a BirA site for single molecule biotinylation was used (49), a generous gift from Dr. Wucherpfenning (Department of Cancer Immunology and AIDS, Dana-Farber Cancer Institute, Boston, MA). TTc was biotinylated by incubation with BirA enzyme at a 1:20 M ratio for 4 h at 30°C in a buffer containing 10 mM Tris) pH 8; 0.1 mM biotin; 10 mM ATP; 10 mM magnesium acetate; and 50 mM bicine, pH 8.3 (49). TTc-QD complexes were prepared as described above in PBS. In control side-by-side experiments using vaccinated adult PBMC, no differences in TT-MBC frequencies were found using either TT-QD or TTc-QD (data not shown).

Thawed lymphocytes were washed with PBS and stained with the amine reactive viability dye ViViD (Invitrogen/Molecular Probes, OR) following the manufacturer’s instructions. Cells were subsequently washed twice in 1× PBS plus 5% FBS, then adjusted to a cell density of 5–10 × 107 cells per ml and incubated with 16 pmol of TT-QD, 20 pmol of DT-QD, 10 pmol of HSA-QD, or 5 pmol of TTc-QD/50 μl stain volume on ice for 30 min with intermittent gentle vortexing. Cells were then costained CD19-allophycocyanin_Cy7 (eBioSciences), CD27PE_Cy7 (BioLegend), and IgD–Alexa 488 (BD Pharmingen, CA) for an additional 30 min on ice. In some experiments with United States PBMC, CD21-allophycocyanin (BD Pharmingen) was added. All Abs were individually titrated. Cells were washed three times with 1.5 ml of PBS/5% FBS and fixed with final concentration of 1% paraformaldehyde in 1× PBS (Electron Microscopy Sciences, CA). The labeled cells were analyzed on a LSRII (BD Biosciences) and data analyzed using FlowJo software (Tree Star). The overall gating strategy is depicted in Supplemental Fig. 1. We used a polychromatic gating strategy to eliminate aberrant binding events. Singlet cells were first selected in an FSC-area (FSC-A) versus FSC-height plot followed by a tight lymphocyte inclusion gated SSC-A versus FSC-A. CD19+ B cells were identified against the ViVid channel to exclude dead cells that could bind Ag-QD complexes and fluorophore-conjugated Abs nonspecifically. Ag-QD+ cells were displayed as CD19+ class-switched MBC (CD19+/CD27+/IgD) events. At least 200,000 viable lymphocytes were gated for all experiments unless otherwise indicated.

Five-day B cell ELISpot assays were performed as previously described (50). Briefly, 1 × 106 PBMC were cultured in complete media in 24-well plates alone or with stimulation consisting of 2.5 μg of CpG oligodeoxynucleotide ODN-2006 (Operon Technologies, CA), 1/10,000 dilution of protein A from Staphylococcus aureus Cowan (Sigma-Aldrich), 1/100,000 dilution of PWM (Sigma-Aldrich), and 25 ng/ml IL-10 (R&D Systems)/ml in a volume of 200 μl for 5 d. Cells were then washed in complete media and plated on prepared ELISpot plates (S2EM004M99; Millipore, MA) previously coated with either PBS, 10 μg of polyclonal goat anti-human IgG (Caltag, CA) to detect all Ab-secreting cells, 2.5 μg of TT, or 4.0 μg of DT/ml. Stimulated PBMC were incubated with Ag for 6 h unless otherwise specified. Plates were developed, scanned, and spots counted by ImmunoSpot satellite analyzer (Cellular Technology, OH). For the ex vivo B cell ELISpot assay, cryopreserved PBMC were thawed, rested, and plated onto plates that were coated with either human anti-IgG or 4 μg DT/ml for 6 h. No stimulation was used.

Data were analyzed using GraphPad Prism 6.0 (GraphPad Software, San Diego, CA). Multiple groups were compared using one- or two-way ANOVA with Bonferroni posttest correction. Two-tailed nonparametric tests analysis with Mann–Whitney and Kruskal–Wallis tests were performed to compare responses between the comparator groups. Comparisons between medians of paired samples were made using Wilcoxon rank test for nonparametric pairs.

Clinical characteristics of the Kenyan mothers, their pregnancy histories, and newborns are shown in Table I. All pregnancies were uncomplicated, and 80% were vaginal deliveries. The comparator groups described include 1) United States cord blood; 2) Kenyan noninfectious exposed cord blood; and 3) Kenyan all infections/infectious exposed cord blood consists of 1) CMV IgM+ cord blood, 2) HIV-exposed cord blood, and 3) P. falciparum–sensitized cord blood. Few differences were detected among the comparator groups with respect to age, sex, gestational age, birth weight, and cord blood hemoglobin levels. Geohelminth infections were uncommon.

FIGURE 1.

Example of gating strategy using United States adult PBMCs. First, singlets are gated followed by lymphocytes in the forward and side scattered gates. Next, live CD19+ cells are gated followed by designation of MBC subsets by differential expression of CD27 and IgD. FSC-H, FSC-height; SSC-A, SSC-area.

FIGURE 1.

Example of gating strategy using United States adult PBMCs. First, singlets are gated followed by lymphocytes in the forward and side scattered gates. Next, live CD19+ cells are gated followed by designation of MBC subsets by differential expression of CD27 and IgD. FSC-H, FSC-height; SSC-A, SSC-area.

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Table I.
Kenyan maternal and neonatal characteristics
K Nonexposed Cord Blood (n = 10)K Infections/Infectious Exposed Cord Blood (n = 25)
AllK CMV+ (n = 4)K HIV-Exposed (n = 13)K P. falciparum–Exposed (n = 8)
Mean maternal age, y (SD) 21 (6) 28 (11) 24 (6) 31 (5) 23 (6) 
Multigravidity, n (%) (≥3 pregnancies) 6 (60) 12 (48) 1 (25) 8 (62) 3 (38) 
Median maternal Hgb, g/dl (range) 7.8 (4.5–11.2) 10.2 (7.4–13.0) 10.7 (9.8–11.6) 9.0 (5.8–10.6) 10.7 (6.6–14.8) 
Maternal parasite, n (%)      
 Hookworm 2 (20) 2 (8) 2 (25) 
Entameba histolytica 3 (30) 3 (12) 3 (38) 
Giardia lamblia 1 (10) 
 Trichuris 
 Ascaris 
Schistosoma hematobium 1 (4) 1 (13) 
 Strongyloides 
Vaginal delivery, n (%) 9 (90) 20 (80) 2 (50) 11 (85)  7 (88) 
Bed net usage, n (%) 3 (30) 17 (68%) 4 (100) 10 (77) 4 (50) 
Median gestational age, wk (range) 38.5 (35.3–41.8) 40 (38.0–42.0) 40.5 (39.5–41.5) 39.0 (36.0–42.0) 39.0 (34.0–44.0) 
Median birth weight, g (range) 2800 (2200–3400) 3000 (2200–3800) 3700 (3600–3700) 3000 (2400–4500) 3000 (1900–4100) 
Male, n (%) 3 (30) 12 (57) 3 (75%) 3 (23%) 7 (88%) 
Median cord blood Hgb, g/dl (range) 14.5 (11.4–17.6) 14.2 (12.2–16.3) 13.3 (12.7–14.0) 14.0 (12.0–16.3) 15.3 (12.3–18.3) 
K Nonexposed Cord Blood (n = 10)K Infections/Infectious Exposed Cord Blood (n = 25)
AllK CMV+ (n = 4)K HIV-Exposed (n = 13)K P. falciparum–Exposed (n = 8)
Mean maternal age, y (SD) 21 (6) 28 (11) 24 (6) 31 (5) 23 (6) 
Multigravidity, n (%) (≥3 pregnancies) 6 (60) 12 (48) 1 (25) 8 (62) 3 (38) 
Median maternal Hgb, g/dl (range) 7.8 (4.5–11.2) 10.2 (7.4–13.0) 10.7 (9.8–11.6) 9.0 (5.8–10.6) 10.7 (6.6–14.8) 
Maternal parasite, n (%)      
 Hookworm 2 (20) 2 (8) 2 (25) 
Entameba histolytica 3 (30) 3 (12) 3 (38) 
Giardia lamblia 1 (10) 
 Trichuris 
 Ascaris 
Schistosoma hematobium 1 (4) 1 (13) 
 Strongyloides 
Vaginal delivery, n (%) 9 (90) 20 (80) 2 (50) 11 (85)  7 (88) 
Bed net usage, n (%) 3 (30) 17 (68%) 4 (100) 10 (77) 4 (50) 
Median gestational age, wk (range) 38.5 (35.3–41.8) 40 (38.0–42.0) 40.5 (39.5–41.5) 39.0 (36.0–42.0) 39.0 (34.0–44.0) 
Median birth weight, g (range) 2800 (2200–3400) 3000 (2200–3800) 3700 (3600–3700) 3000 (2400–4500) 3000 (1900–4100) 
Male, n (%) 3 (30) 12 (57) 3 (75%) 3 (23%) 7 (88%) 
Median cord blood Hgb, g/dl (range) 14.5 (11.4–17.6) 14.2 (12.2–16.3) 13.3 (12.7–14.0) 14.0 (12.0–16.3) 15.3 (12.3–18.3) 

All p values > 0.1.

Hgb, hemoglobin; K, Kenyan.

To examine the impact of CMV, HIV, and/or malaria infection during pregnancy on B cell development and maturation in the fetus, we measured plasma cytokines, some of which are associated with B cell activation and differentiation, in cord blood of various comparator groups. We compared the United States cord blood to the Kenyan noninfectious exposed cord blood and found that Kenyan neonates had statistically significant higher levels of IP-10, TNF-α, CRP, sCD14, sTNFR1, sTNFR2, and BAFF (Fig. 2). United States cord blood had slightly higher levels of IL-7, IL-12 (p40), IL-12 (p70), and IL-17a compared with the Kenyan noninfectious exposed cord blood. No statistically significant differences among the groups were observed for IL-4, IL-6, IL-9, or IL-10. Next, we compared United States cord blood to the Kenyan all infections/infectious exposure group and found that the Kenyan neonates had higher levels of IP-10, TNF-α, CRP, sCD14, sTNFR1, and BAFF compared with United States cord blood. Within the Kenyan cord blood groups, those with documented infections/infectious exposures had statistically significant higher levels of IFN-γ, IL-7, and IL-17a compared with the noninfectious exposed group (Fig. 2). Although Kenyan cord blood had higher CRP values than United States cord blood, the values for all groups were within a normal range. Thus, most proinflammatory biomarkers, including those associated with B cell development and maturation, were higher in Kenyan cord blood groups than United States cord blood, especially among newborns of mothers with HIV or malaria or born with congenital CMV.

FIGURE 2.

Plasma cytokines/chemokines in cord blood of comparator groups (bottom x-axes). United States cord blood (solid circles), Kenyan noninfectious exposed cord blood (solid triangles), Kenyan all infections/infectious exposure group cord blood (K all inf) (solid diamonds), Kenyan CMV+ cord blood (open diamonds), Kenyan HIV-exposed cord blood (open circles), and Kenyan P. falciparum–exposed cord blood (open squares). Medians shown with comparisons made by Mann–Whitney tests. All measurements were in picograms per milliliter except for sCD14 and CRP, which are measured in nanograms per milliliter as indicated. Some values were log transformed for ease of visualization. The p values ≤ 0.05 are indicated. exp, exposed; K, Kenyan.

FIGURE 2.

Plasma cytokines/chemokines in cord blood of comparator groups (bottom x-axes). United States cord blood (solid circles), Kenyan noninfectious exposed cord blood (solid triangles), Kenyan all infections/infectious exposure group cord blood (K all inf) (solid diamonds), Kenyan CMV+ cord blood (open diamonds), Kenyan HIV-exposed cord blood (open circles), and Kenyan P. falciparum–exposed cord blood (open squares). Medians shown with comparisons made by Mann–Whitney tests. All measurements were in picograms per milliliter except for sCD14 and CRP, which are measured in nanograms per milliliter as indicated. Some values were log transformed for ease of visualization. The p values ≤ 0.05 are indicated. exp, exposed; K, Kenyan.

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Although cytokines do not cross the placenta, we compared cytokine levels in Kenyan maternal plasma (at delivery) to paired cord blood plasma with the expectation that mothers would have higher levels of inflammation than their neonate but found few differences. The exceptions to this were BAFF levels in maternal–neonate pairs and cytokines in the P. falciparum–exposed groups. Fig. 3 illustrates BAFF differences between maternal–neonate pairs. Essentially, median levels of BAFF in Kenyan cord blood (noninfectious exposed and all infection/infectious exposed) were double that of their paired maternal plasma. When comparing the P. falciparum–exposed cord blood to their maternal samples, cord blood contained higher levels of IP-10, sTNFR1, TNF-α, IL-7, and IL-12 (p70) (Fig. 4).

FIGURE 3.

Plasma BAFF levels in Kenyan maternal–neonatal pairs. Plasma BAFF levels (picograms per milliliter) log transformed were compared between maternal and neonate pairs using Wilcoxon matched pairs test; medians shown. The comparator groups (x-axis) include noninfectious exposed maternal (solid upside down triangles), noninfectious exposed cord blood (open upside down triangles), all infection/infectious exposed maternal (solid triangles), all infections/infectious exposed cord blood (open triangles), mothers of neonates with congenital CMV (Mat CMV) (solid diamonds), CMV+ neonatal cord blood (open diamonds), HIV+ maternal (solid circles), HIV-exposed cord blood (open circles), P. falciparum+ maternal (solid squares), and P. falciparum–exposed cord blood (open squares). Medians shown with comparisons made by Wilcoxon rank test. The p values ≤0.05 are indicated. exp, exposed; Mat, maternal; Neo, neonatal.

FIGURE 3.

Plasma BAFF levels in Kenyan maternal–neonatal pairs. Plasma BAFF levels (picograms per milliliter) log transformed were compared between maternal and neonate pairs using Wilcoxon matched pairs test; medians shown. The comparator groups (x-axis) include noninfectious exposed maternal (solid upside down triangles), noninfectious exposed cord blood (open upside down triangles), all infection/infectious exposed maternal (solid triangles), all infections/infectious exposed cord blood (open triangles), mothers of neonates with congenital CMV (Mat CMV) (solid diamonds), CMV+ neonatal cord blood (open diamonds), HIV+ maternal (solid circles), HIV-exposed cord blood (open circles), P. falciparum+ maternal (solid squares), and P. falciparum–exposed cord blood (open squares). Medians shown with comparisons made by Wilcoxon rank test. The p values ≤0.05 are indicated. exp, exposed; Mat, maternal; Neo, neonatal.

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FIGURE 4.

Maternal–neonatal plasma cytokines in P. falciparum–sensitized pairs. Solid squares represent maternal plasma cytokine levels at delivery, and open squares represent P. falciparum–sensitized neonatal cord blood cytokine levels. Cytokines examined are labeled on the x-axis of each pair. All cytokines were measured in picograms per milliliter. Medians shown. The p values for differences are above the pairs. Differences were detected using Wilcoxon rank test.

FIGURE 4.

Maternal–neonatal plasma cytokines in P. falciparum–sensitized pairs. Solid squares represent maternal plasma cytokine levels at delivery, and open squares represent P. falciparum–sensitized neonatal cord blood cytokine levels. Cytokines examined are labeled on the x-axis of each pair. All cytokines were measured in picograms per milliliter. Medians shown. The p values for differences are above the pairs. Differences were detected using Wilcoxon rank test.

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The frequency of B cells were similar among the groups. Kenyan cord blood contained higher frequencies of CD4 T cells (median 76.9% Kenyan nonexposed, 73% Kenyan all infections/infectious exposure group) and lower frequencies of CD8 T cells (median 20.6% Kenyan nonexposed, 20.9% Kenyan all infections/infectious exposure group) compared with United States cord blood (median 55.8 and 40.9% respectively, Fig. 5). The frequency of B cells, CD4 T cells, or CD8 T cells were similar in cord blood from Kenyan nonexposed and infection/infectious exposure groups (Fig. 5). For determination of T cell frequencies, four HIV-exposed, uninfected CBMC samples did not have enough cells to perform this flow analysis, as our priority was examination of B cell subset frequencies.

FIGURE 5.

B cell, CD4, and CD8 T cell frequencies in the cord blood comparator groups. The x-axis is labeled with cord blood comparator groups including United States cord blood (solid circles), Kenyan noninfectious exposed cord blood (solid triangles), Kenyan all infections/infectious exposure group cord blood (solid diamonds), Kenyan CMV+ cord blood (open diamonds), Kenyan HIV-exposed cord blood (open circles), and Kenyan P. falciparum–exposed cord blood (open squares). Medians shown with comparisons made by Mann–Whitney tests. The p values ≤0.05 are indicated. exp, exposed; K, Kenyan.

FIGURE 5.

B cell, CD4, and CD8 T cell frequencies in the cord blood comparator groups. The x-axis is labeled with cord blood comparator groups including United States cord blood (solid circles), Kenyan noninfectious exposed cord blood (solid triangles), Kenyan all infections/infectious exposure group cord blood (solid diamonds), Kenyan CMV+ cord blood (open diamonds), Kenyan HIV-exposed cord blood (open circles), and Kenyan P. falciparum–exposed cord blood (open squares). Medians shown with comparisons made by Mann–Whitney tests. The p values ≤0.05 are indicated. exp, exposed; K, Kenyan.

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In regard to the B cell subsets, no differences among the United States and Kenyan cord blood comparator groups were detected for mature, immature, and transitional B cell populations (Supplemental Fig. 2). We next focused on MBC subsets (Fig. 6). United States cord blood had higher classic isotype-switched MBC frequencies (median 1.04%) than Kenyan nonexposed cord blood (median 0.39%, p = 0.0167). United States cord blood also had higher proportions of non–class-switched MBC (median 1.69%) compared with Kenyan noninfectious exposed and Kenyan all infections/infectious exposure group frequencies (median 0.95%, p = 0.0085, median 0.73, p = 0.0015 respectively). Frequencies of classic and nonswitched MBC were low in all cord blood comparator groups (<3% of B cells). For activated MBC, both Kenyan noninfectious (median 2.74%) and Kenyan all infections/infectious exposure group (median 3.25%) cord blood had higher frequencies than United States cord blood (median 0.51%, p < 0.001 and p = 0.001 respectively). Additionally, the Kenyan all infections/infectious exposure group had higher proportions of atypical MBC (median 7.77%) compared with United States cord blood (median 4.73%, p = 0.0103). Interestingly, this was driven by the Kenyan HIV-exposed group that had the highest frequencies of atypical MBC (median 13.39%).

FIGURE 6.

MBC subset frequencies among the cord blood comparator groups. The x-axis is labeled with cord blood comparator groups including United States cord blood (solid circles), Kenyan noninfectious exposed cord blood (solid triangles), Kenyan all infections/infectious exposure group cord blood (solid diamonds), Kenyan CMV+ cord blood (open diamonds), Kenyan HIV-exposed cord blood (open circles), and Kenyan P. falciparum–exposed cord blood (open squares). Medians shown with comparisons made by Mann–Whitney tests. The p values ≤0.05 are indicated. exp, exposed; K, Kenyan.

FIGURE 6.

MBC subset frequencies among the cord blood comparator groups. The x-axis is labeled with cord blood comparator groups including United States cord blood (solid circles), Kenyan noninfectious exposed cord blood (solid triangles), Kenyan all infections/infectious exposure group cord blood (solid diamonds), Kenyan CMV+ cord blood (open diamonds), Kenyan HIV-exposed cord blood (open circles), and Kenyan P. falciparum–exposed cord blood (open squares). Medians shown with comparisons made by Mann–Whitney tests. The p values ≤0.05 are indicated. exp, exposed; K, Kenyan.

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We specifically examined IgM+CD27+ MBC, as they represent an interesting population whose nature and origin are debated. They are thought to have exited the germinal center early, act in a T cell–independent manner, may have immunoregulatory properties, and the Abs they produce could act as a rapid first-line defense for the neonate (51, 52). Some studies contend that cord blood does not contain IgM+ MBC (53), whereas others found that cord blood contains ∼2% IgM+CD27+ MBC (54). We found that United States cord blood had ∼2% IgM+CD27+ MBC. Among the cord blood comparator groups, United States cord blood had a higher frequency of CD27+IgM+IgD+ MBC (median 1.69%) than Kenyan nonexposed (median 0.95%, p = 0.0085) and Kenyan all infections/infectious exposure group cord blood (median 0.75%, p = 0.0049). Kenyan HIV-exposed cord blood had slightly higher CD27+IgM+IgD MBC frequencies than Kenyan nonexposed cord blood (median 0.4% versus 0.21% of B cells, p = 0.0234). Otherwise, the IgM+CD27+ MBC populations were small and comparable among the cord blood groups (Fig. 7).

FIGURE 7.

IgM+ MBC subset frequencies among the cord blood comparator groups. The x-axis is labeled with cord blood comparator groups including United States cord blood (solid circles), Kenyan noninfectious exposed cord blood (solid triangles), Kenyan all infections/infectious exposure group cord blood (K all inf) (solid diamonds), Kenyan CMV+ cord blood (open diamonds), Kenyan HIV-exposed cord blood (open circles), and Kenyan P. falciparum–exposed cord blood (open squares). Medians shown with comparisons made by Mann–Whitney tests. The p values ≤0.05 are indicated. exp, exposed; K, Kenyan.

FIGURE 7.

IgM+ MBC subset frequencies among the cord blood comparator groups. The x-axis is labeled with cord blood comparator groups including United States cord blood (solid circles), Kenyan noninfectious exposed cord blood (solid triangles), Kenyan all infections/infectious exposure group cord blood (K all inf) (solid diamonds), Kenyan CMV+ cord blood (open diamonds), Kenyan HIV-exposed cord blood (open circles), and Kenyan P. falciparum–exposed cord blood (open squares). Medians shown with comparisons made by Mann–Whitney tests. The p values ≤0.05 are indicated. exp, exposed; K, Kenyan.

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In summary, Kenyan cord blood had an altered distribution of MBC subpopulations compared with United States cord blood with lower but detectable IgM+ MBC, more activated MBC, and elevated proportions of atypical MBC among the Kenyan HIV-exposed cord blood samples.

CD5 on B cells can arise naturally or be induced by stimuli (55). CD5 modulates B cell activity and may have a role in innate immune responses and B cell regulation and survival through the production of IL-10 (56, 57). TLR2 expression in B cells is important for both innate and adaptive functions. Neonatal B cell TLR2-mediated activity from CBMC from developed countries is impaired compared with adult B cells (58). We hypothesized that Kenyan fetuses may have a higher Ag exposure and therefore have higher surface CD5 and TLR2 expression and differential responses to stimuli.

Ex vivo, we measured the proportion of B cells expressing CD5 and TLR2. Previous reports found that the cord blood B cell compartment contains ∼60% CD5+ B cells that decrease with age such that adults have ∼5–30% CD5+ B cells (59, 60). This proportion may be variable, as Köhler et al. (61) found CD5+ B cell frequencies to be ∼10% in Gabonese CBMC and ∼15% in Austrian CBMC. Kenyan noninfectious exposed cord blood had comparable CD5+ B cell frequencies to United States cord blood. The Kenyan all infections/infectious exposure group had higher frequencies of CD5+ B cells (median 10.5%) compared with the Kenyan noninfectious exposed group (median 6.53%, p = 0.0272) and United States cord blood (median 5.57%, p = 0.003, Fig. 8). No statistically significant differences were detected among the infectious exposure cord blood subgroups. Examining CD5 expression in CD27+ MBC, we found that the Kenyan all infections/infectious exposure group had higher proportions (median 8.84%) compared with the Kenyan noninfectious exposed group (median 5.77%, p = 0.024) and United States cord blood (median 2.467%, p < 0.001, Fig. 8). No statistically significant differences were detected among the infections/infectious exposure subgroups. B cell surface expression of TLR2 ranged between medians of 1.4–2.2%, with no statistical differences detected among all the comparator groups. Of the CD27+ MBC, TLR2+ frequencies were very low and ranged from 0.38 to 0.51% with no differences detected among the cord blood comparator groups (data not shown).

FIGURE 8.

CD5+ B cell frequencies among the cord blood comparator groups. Top panel shows CD5+ B cell frequencies. Bottom panel shows CD5+ CD27+ B cells. The x-axis is labeled with cord blood comparator groups including United States cord blood (solid circles), Kenyan noninfectious exposed cord blood (solid triangles), Kenyan all infections/infectious exposure group cord blood (K all inf) (solid diamonds), Kenyan CMV+ cord blood (open diamonds), Kenyan HIV-exposed cord blood (open circles), and Kenyan P. falciparum–exposed cord blood (open squares). Medians shown with comparisons made by Mann–Whitney tests. The p values ≥ 0.05 are indicated. exp, exposed; K, Kenyan.

FIGURE 8.

CD5+ B cell frequencies among the cord blood comparator groups. Top panel shows CD5+ B cell frequencies. Bottom panel shows CD5+ CD27+ B cells. The x-axis is labeled with cord blood comparator groups including United States cord blood (solid circles), Kenyan noninfectious exposed cord blood (solid triangles), Kenyan all infections/infectious exposure group cord blood (K all inf) (solid diamonds), Kenyan CMV+ cord blood (open diamonds), Kenyan HIV-exposed cord blood (open circles), and Kenyan P. falciparum–exposed cord blood (open squares). Medians shown with comparisons made by Mann–Whitney tests. The p values ≥ 0.05 are indicated. exp, exposed; K, Kenyan.

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To examine the differential responsiveness of B cells with different in utero infectious exposures to stimuli, we cultured CBMC with or without polyclonal stimulation mixture for 12 h and examined the expression of CD5 and TLR2 in B cell subsets. In general, as observed by others (62), we found there was considerable spontaneous B cell death. However, between the mononuclear cells with and without stimuli, there was no difference in the proportion of B cells after 12 h (e.g., stimuli did not promote cell death). We found that after 12 h, the Kenyan P. falciparum–exposed group-stimulated cells had higher levels of CD5+ B cells (median 47.85%) compared with those without stimulus (median 23.9%, p = 0.0078, Fig. 9). This drove the observation that the Kenyan all infections/infectious exposure group with stimulus had higher frequencies of CD5+ B cells (median 54.65%) compared with the nonstimulated cells (median 35.35%, p = 0.0005). The majority of the CD5+ B cells were also CD27+, and the same response to stimuli was observed (Fig. 9). TLR2 expression on B cells did not change significantly with stimulus for any of the comparator groups and remained low at ∼2–5% for all groups (data not shown).

FIGURE 9.

CD5+ B cell frequencies after 12 h of culture with media or polyclonal stimulation. Top panel shows CD5+ B cell frequencies in the cord blood comparator groups. Solid symbols represent culture with media only. Open symbols represent cells from the same group after culture with stimulation. Bottom panel shows CD5+ CD27+ B cells. Medians shown. exp, exposed; K, Kenyan; stim, stimulation.

FIGURE 9.

CD5+ B cell frequencies after 12 h of culture with media or polyclonal stimulation. Top panel shows CD5+ B cell frequencies in the cord blood comparator groups. Solid symbols represent culture with media only. Open symbols represent cells from the same group after culture with stimulation. Bottom panel shows CD5+ CD27+ B cells. Medians shown. exp, exposed; K, Kenyan; stim, stimulation.

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Having observed changes in B cell memory subpopulations and cytokine levels in neonates whose mothers had infectious in utero exposures, we sought to understand if this affected fetal priming to specific Ags. Kenyan women in their first, second, and third pregnancies receive TT vaccinations to prevent neonatal tetanus. HIV+ women are known to have lower titers of TT despite vaccinations (63). Because HIV-exposed neonates had elevated atypical MBC, we hypothesized that they may have compromised capacity to make Ag-specific classic MBC (i.e., TT-MBC) compared with non–HIV-exposed neonates. We developed a flow-based assay that uses Ags bound to QD to detect low-frequency TT-MBC frequencies in cord blood. To validate the assay, we examined the dynamics of DT-MBC and TT-MBC frequencies in healthy United States adults who received the tetanus, diphtheria, and acellular pertussis vaccine. We examined Ag-specific MBC frequencies in PBMC at day 0, 7, and 15 by ELISpot and flow cytometry. Supplemental Fig. 1 depicts the gating strategy to determine Ag-specific classic MBC frequencies in thawed PBMC. We compared DT-MBC frequencies by ex vivo ELISpot (which detects DT-specific plasma cells [PC]), 5-d polyclonal ELISpot (which detects DT-MBC), and ex vivo flow assay (detects both DT-PC and DT-MBC) (Supplemental Fig. 3). We found that the ex vivo B cell ELISpot detected DT-specific PC that reached their peak frequency in peripheral blood at 7 d postvaccination and were no longer detected by day 15. In contrast, the 5-d stimulated B cell ELISpot detected DT-specific Ab-secreting cells and the flow assay–measured DT-MBC frequencies peaked at day 15. TT-MBC frequencies were determined at the same time and in five previously vaccinated United States adult donors with boosting TT-MBC frequencies by day 15 (Supplemental Fig. 4). The dynamics of Ag-specific MBC formation after vaccination is consistent with other studies showing a boosting of Ag-specific MBC at 15 d postvaccination (64). Additionally, the frequency of TT- and DT-MBC using this flow assay is comparable to that observed with other flow-based Ag detection assays (50, 65).

We used this flow-based assay to evaluate the frequency of TTc-specific MBC in United States cord blood (n = 16) and Kenyan cord blood (n = 52). In the Kenyan cord blood cohort, 26 were born to HIV+ mothers and 26 were born to HIV− mothers. Characteristics of this cohort are described in Table II. In general, the groups were similar except that HIV+ mothers were slightly older than HIV− mothers (mean 29 y versus 24 y, p = 0.0018) and had slightly heavier neonates (median 3400 g versus 3200 g, p = 0.0267). A very small number of worm infections were detected in HIV− mothers. Compared with the validation experiments described previously, greater numbers of viable lymphocytes were examined (∼4 × 105) to detect TTc-MBC rare events. An average of 4.01 × 105 lymphocytes from United States CBMC and 4.23 × 105 lymphocytes from Kenyan CBMC were acquired. No statistical differences between lymphocyte numbers, B cell numbers, or B cell frequencies were detected among the groups. Thus, differences in detected TT-MBC frequencies were not a function of different lymphocyte or B cell numbers acquired. In the data presented in this paper, one TT-MBC (if detected) was subtracted from the TT-MBC counts of each individual to set a minimal threshold and account for background noise. Data are presented as absolute TT-MBC counts as well as TT-MBC frequencies of the classic MBC population. The rationale for data presentation in this manner is that cord blood contains few classic MBC, and thus TT-MBC frequencies are affected by the absolute number of MBC detected. We found that United States cord blood had lower TT-MBC counts (median 0) compared with Kenyan non–HIV-exposed (median 2, p = 0.0004) and Kenyan HIV-exposed (median 4.5, p < 0.0001) cord blood. Kenyan HIV-exposed TT-MBC counts were slightly higher than Kenyan non-HIV TT-MBC counts (Fig. 10A). There was no difference in TT-MBC frequencies between the Kenyan groups, but both had higher proportions of TT-MBC frequencies (median 2.13 and 3.15%, respectively) compared with United States cord blood (Fig. 10B). All but two Kenyan women were experiencing their first, second, or third pregnancy and received tetanus booster vaccinations as recorded in their medical records. Of the 16 United States CBMC samples, only three had detectable TT-MBC (range 1–2 events). These samples were obtained prior to the recommendation in the United States to vaccinate United States pregnant women in the second trimester with tetanus, diphtheria, and acellular pertussis vaccine because of community-waning pertussis immunity. HIV-exposed neonates had at least as good a response to TT vaccine given to mothers during pregnancy as non–HIV-exposed Kenyan neonates based on their TT-MBC frequencies.

Table II.
Kenyan HIV+ and HIV− maternal and neonatal characterizations
HIV+ (n = 26)HIV− (n = 26)p Value
Mean maternal age, y (SD) 29 (5) 24 (4) 0.0018 
Multigravidity, n (%) (≥3 pregnancies) 12 (46) 6 (23)  
Median maternal Hgb at delivery, g/dl (range) 10.4 (5.8–13.2) 10.2 (5.2–12.9)  
Median CD4 count at enrollment (range) 425 (157–1100) Not done  
Maternal parasite, n (%)    
 Hookworm 3 (12) 
 Entameba histolytica 2 (8) 
Giardia lamblia 1 (4) 
Trichuris 
 Ascaris 
Schistosoma hematobium 2 (8) 
 Strongyloides 
Vaginal delivery, n (%) 24 (92) 24 (92)  
Bed net usage, n (%) 24 (92) 22 (85)  
Median gestational age, wk (range) 39 (36.5–41) 40 (38.0–42.0)  
Median birth weight, g (range) 3400 (2500–4500) 3200 (2400–4200) 0.0267 
Male, n (%) 10 (38) 16 (62)  
Median cord blood Hgb, g/dl (range) 13.4 (11.8–16) 14.1 (12.2–17.6)  
HIV+ (n = 26)HIV− (n = 26)p Value
Mean maternal age, y (SD) 29 (5) 24 (4) 0.0018 
Multigravidity, n (%) (≥3 pregnancies) 12 (46) 6 (23)  
Median maternal Hgb at delivery, g/dl (range) 10.4 (5.8–13.2) 10.2 (5.2–12.9)  
Median CD4 count at enrollment (range) 425 (157–1100) Not done  
Maternal parasite, n (%)    
 Hookworm 3 (12) 
 Entameba histolytica 2 (8) 
Giardia lamblia 1 (4) 
Trichuris 
 Ascaris 
Schistosoma hematobium 2 (8) 
 Strongyloides 
Vaginal delivery, n (%) 24 (92) 24 (92)  
Bed net usage, n (%) 24 (92) 22 (85)  
Median gestational age, wk (range) 39 (36.5–41) 40 (38.0–42.0)  
Median birth weight, g (range) 3400 (2500–4500) 3200 (2400–4200) 0.0267 
Male, n (%) 10 (38) 16 (62)  
Median cord blood Hgb, g/dl (range) 13.4 (11.8–16) 14.1 (12.2–17.6)  

Hgb, hemoglobin.

FIGURE 10.

TT-specific MBC in United States cord blood compared with Kenyan non–HIV-exposed and HIV-exposed cord blood. (A) shows the absolute counts of TT-specific classic MBC in United States cord blood, Kenyan non–HIV-exposed cord blood, and Kenyan HIV-exposed cord blood. (B) shows TT-specific MBC as a frequency of the total classic MBC population for the cord blood comparator groups. Medians shown. exp, exposed; K, Kenyan.

FIGURE 10.

TT-specific MBC in United States cord blood compared with Kenyan non–HIV-exposed and HIV-exposed cord blood. (A) shows the absolute counts of TT-specific classic MBC in United States cord blood, Kenyan non–HIV-exposed cord blood, and Kenyan HIV-exposed cord blood. (B) shows TT-specific MBC as a frequency of the total classic MBC population for the cord blood comparator groups. Medians shown. exp, exposed; K, Kenyan.

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We sought to examine how infections during successful term pregnancy affect fetal B cell development. To this end, we examined cord blood proinflammatory biomarkers and B cell subset frequencies in Kenyan neonates with congenital CMV and neonates with in utero exposures to HIV or P. falciparum or no infectious exposures. We found that 1) Kenyan neonates, especially those with or exposed to in utero infections, had elevated levels of plasma proinflammatory cytokines compared with United States neonates, 2) Kenyan cord blood plasma BAFF levels were higher than paired maternal levels, 3) Kenyan neonates had shifts in MBC subsets with overall increased frequencies of activated and atypical MBC compared with United States cord blood, 4) Kenyan infection/infectious exposed neonates had higher frequencies of CD5+ B cells, which expanded after stimulation compared with Kenyan neonates with no identified in utero infectious exposures, and 5) Kenyan HIV-exposed neonates exhibited equivalent capacity to generate TTc-specific MBC as non–HIV-exposed neonates despite shifts in MBC subset distributions. Uniquely, P. falciparum–sensitized neonates had several elevated cord blood plasma cytokine/chemokines compared with their paired maternal plasma levels, and HIV-exposed neonates had increased proportions of atypical MBC.

Cytokine production during pregnancy is altered by microbial infections, and proinflammatory cytokines such as IL-6 and TNF-α have been associated with adverse events during pregnancy (6669). Elevated intervillous placental cytokines/chemokines, especially IP-10, were associated with HIV mother-to-child transmission in Malawi (70). Cord blood IP-10 levels of Kenyan neonates in our study were similar to the lower placental IP-10 levels of Malawian HIV-infected nontransmitter pairs (cord blood cytokines were not measured in this study). Multiple studies have shown that P. falciparum infection in pregnancy is associated with adverse outcomes and elevated proinflammatory cytokines/chemokines (7175). Few, however, have shown differences between maternal plasma levels at delivery and cord blood levels. Djontu et al. (76) showed that Cameroonian women with P. falciparum during pregnancy had higher intervillous placental plasma levels of IL-28A and IL-27, but maternal peripheral and cord blood levels were comparable. In this study, for the first time, to our knowledge, we found that P. falciparum–exposed neonates had several elevated biomarkers (IP-10, sTNFR1, TNF-α, IL-7, and IL-12 [p70]) compared with their paired maternal plasma levels. The P. falciparum–exposed group may represent a bias in that we determined P. falciparum exposure not just by the presence of P. falciparum in the maternal blood but also by CBMC recognition and response to P. falciparum Ags, which were transported across the placenta to the fetus, resulting in immune priming. Multiple P. falciparum Ags and perhaps those of nonmalaria origins presumably stimulated this inflammatory response. BAFF, a cytokine member of the TNF family, is an important cytokine for maturation of immature B cells (62, 7779). We found that Kenyan neonates had higher BAFF levels than United States neonates, and Kenyan neonates had higher BAFF levels than their mothers at delivery, a finding also observed by others (80, 81). Lundell et al. (82) found BAFF levels were maximal at birth, higher in cord blood than paired maternal samples, but also higher in neonates born to farming mothers (with presumably a higher Ag exposure) compared with nonfarming cord blood. BAFF levels have also been observed to be elevated in children with acute malaria (83).

Several studies have included the distribution of B cells in their characterization of lymphocytes in cord blood (8486). Two separate studies from Poland and the Netherlands have established reference values of different B cell subsets from different pediatric age groups, including cord blood (54, 87). The total B cell frequencies in the various comparator groups of our study were noted to be lower compared with the Piatosa study (54) but similar to the van Gent study (87). Köhler et al. (61) examined the differences in the phenotypes of B cell, T cell, and monocyte components of CBMC in neonates born to Austrian and Gabonese mothers. No significant differences in the proportion of CD19+ B cells was observed between the two groups even when the Gabonese group was stratified according to malaria infection status, similar to our observations.

The shift in the distribution of MBC subsets in the Kenyan neonates compared with United States neonates likely reflects the differential exposure to environmental Ags, including pathogens and inflammation, resulting in the increase in the proportion of the activated MBC in the Kenyan cord blood group. Interestingly, HIV-exposed neonates had elevated frequencies of atypical MBC. Expansion of this subpopulation of MBC has been described in PBMC of infants as well as adults with HIV or P. falciparum malaria infections (30, 31, 88). In Kenya, HIV-infected children with high viremia had increased proportions of activated MBC and atypical MBC compared with community controls and children with low viremia (89). Recently, a study in Brazil compared plasma cytokine profiles and immunological cell subsets in HIV-exposed and non–HIV-exposed children from birth to 6–12 y of age. At birth, no differences in cytokine levels, B cells, or MBC subsets (including atypical MBC) were found between the groups (90). Finally, pregnant women in Papua New Guinea with high malaria exposure also had elevated frequencies of atypical MBC, but cord blood MBC subset distribution was not examined (91). Thus, HIV or malaria infections are associated with expansion of activated and atypical MBC as we also observed in this study.

It is unclear if inflammation or Ag-induced changes in B cell maturation manifesting as shifts of MBC subset frequencies also affects B cell function. We approached this in two ways by measuring 1) cord blood B cell viability and responses to polyclonal stimulation and 2) fetal priming and formation of Ag-specific MBC formation in response to maternal vaccination. Neonatal CBMC are generally considered immature, but several studies have shown that various cell types are responsive to stimuli. Polyclonal stimulation of CBMC and adult PBMC resulted in expansion of the T regulatory population with those derived from CBMC having stronger immunosuppressive capacities (92). Ex vivo TLR2 stimulation of neonatal naive CD4 T cells resulted in robust responses equivalent to adult naive CD4 T cells (93). In neonates exposed to infections in utero, CD4 and CD8 T cells from hepatitis C virus–exposed neonates had higher IFN-γ production in response to polyclonal stimulation than did T cells from controls (94). Cord blood B cells from noninfectious exposed neonates can produce IgM and some IgG in response to CpG oligodeoxynucleotides but require between 3 and 15 d of stimulation (95). We found that temporally short polyclonal stimulation resulted in significant B cell apoptosis but did not differ significantly among the groups. Infectious exposed neonates retained a higher frequency of CD5+ B cells after stimulation compared with noninfectious exposed neonates. The effect of infectious exposure on the proportion of CD5 in CD19+ B lymphocytes in CBMC has been variable in the literature. Borges-Almeida et al. examined the impact of HIV infection and highly active antiretroviral treatment on neonatal B lymphocyte maturation in full-term HIV-exposed but uninfected neonates. They found that cord blood B cells were increased in the HIV-exposed neonates, and this was mainly driven by an increase in CD5+ B cells (96). Conversely, studies in Gabon have described a lower proportion of CD5+ B lymphocyte in the Gabonese cord blood with evidence of prenatal infectious exposure (e.g., malaria or schistosomiasis) compared with European controls (61, 97). Kessel et al. (62) found that cord blood B cells underwent apoptosis at a higher frequency than adult B cells but that CD5+ B cells from cord blood or adults had a lower rate of apoptosis compared with CD5 B cells. We also observed that the proportion of cord blood CD5+ B cells that were CD27+ were greater in the Kenya all infections group than the United States cord blood (∼10 versus 2%). Although there are, to our knowledge, no comparable data from studies of cord blood B cells in newborns with or without in utero infection exposure, Zhang et al. (98) reported that 10% of CD5+ B cells in peripheral blood of adults with active tuberculosis were CD27+. Collectively, these data indicate that infections and infectious exposures in utero may promote CD5+ B cell development, contributing to improved B cell survival and maturation. Future studies of cord blood CD5+ B cells will need more detailed characterization of their role in regulating immune function, particularly their capacity to produce IL-10 in the context of inflammation resulting from maternal infection (99).

Evidence for in utero sensitization or priming to various Ags has been observed by several groups (e.g., TT [cord blood IgM detection] [100], malaria proteins [cord blood T cell recall responses] (11, 38), helminth proteins [cord blood T cell recall responses] (101), schistosome and filarial Ags [cord blood IgM and IgE detection] (97, 102), and HIV [cord blood T cell recall responses] (5). Some studies have indicated that fetal recall responses to foreign Ags at birth were associated with subsequent increased infant immune responses to these infections and protection from infection compared with infants with no recall responses at birth (10, 21). In a study by Gill et al. (100), babies born of mothers immunized with TT vaccine during pregnancy had TT-specific IgM Ab in cord blood as well as a faster response to TT vaccination during infancy, likely due to sensitization of the fetus in utero. Similar results were shown in other vaccination studies of pregnant mothers using meningococcal and influenza vaccines, where Ag-specific Ab and T cell responses were elicited from the cord blood (103, 104). Thus, the fetal immune system is capable of making appropriate and possibly beneficial immune responses. This is the first report, to our knowledge, using flow cytometry to identify fetal TTc-specific MBC in response to TT vaccination during pregnancy. HIV exposure did not compromise the formation of TT-MBC despite this group having more activated and atypical MBC. It is not known whether these TT-primed infants have higher TT IgG after standard infant vaccinations compared with non–TT-primed infants. These results suggest infectious exposed neonates have preserved B cell function despite MBC subset shifts and experience with proinflammatory cytokines/chemokines. Indeed, several studies show that HIV-exposed uninfected infants have appropriate vaccine responses to immunizations that are comparable to non–HIV-exposed infants (105109).

This study has some limitations that may influence the results. First, as previously mentioned, bias may have been introduced by examining P. falciparum–sensitized neonates rather than those whose mothers only had P. falciparum+ blood samples during pregnancy. Additionally, the CMV+ group was defined by high titers of cord blood CMV IgM rather than the gold standard of neonatal CMV Ag or PCR detection in urine, which was not feasible at the time of the study. We did not examine infectious disease Ag-specific MBC such as P. falciparum–specific MBC, which have been performed by Lugaajju et al. (110) and show P. falciparum–specific MBC in various subsets. Also, inclusion of the two neonates with congenital CMV and exposed to HIV into the HIV-exposed group could have influenced the results. However, when measurements from these two neonates were removed, the final results were not significantly altered. Lack of maternal immunologic characterization was not possible and could have influenced the interpretation of the results. Finally, the relatively small sample sizes may have limited power to detect statistically significant differences among the comparator groups.

Our findings indicate that infections during pregnancy can lead to fetal immune activation affecting B cell development but maintaining humoral immune function. Infections during pregnancy are often deleterious, but in some cases, fetal Ag exposure could prime the immune system such that early infant responses to infectious challenges may be augmented. The long-term impact of these changes needs to be further evaluated to determine if there are any potential implications in the response to infections and vaccinations during infancy.

We thank the study participants and Dr. Penny Holding for overseeing study site management and sample collection in Kenya.

This work was supported by the Center for AIDS Research services of the National Institutes of Health (NIH) P30 AI036219, MH080601 (to Penny Holding), AI064667 (to C.K.), and AI098511 (to A.D.). This publication was made possible by the Clinical and Translational Science Collaborative of Cleveland, Case Western Reserve University/Cleveland Clinic Clinical Translational Science Award UL1TR000439 from the National Center for Advancing Translational Sciences component of the NIH, and NIH Roadmap for Medical Research. Its contents are solely the responsibility of the authors and do not necessarily represent the official views of the NIH.

The online version of this article contains supplemental material.

Abbreviations used in this article:

CBMC

cord blood mononuclear cell

DT

diphtheria toxin

FSC

forward scatter

FSC-A

FSC-area

HSA

human serum albumin

MBC

memory B cell

MFI

mean fluorescence intensity

PC

plasma cell

QD

quantum dot

SSC

side scatter

TT

tetanus toxoid

TTc

TT C-fragment.

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The authors have no financial conflicts of interest.

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