Intrathymic differentiation of natural antibody-producing plasma cells in human neonates

doi:10.21203/rs.3.rs-58978/v1

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Intrathymic differentiation of natural antibody-producing plasma cells in human neonates

https://doi.org/10.21203/rs.3.rs-58978/v1

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published 01 Oct, 2021

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The thymus is a central lymphoid organ responsible for the development of T cells. Here, we show that the thymus of human neonates also contains a consistent contingent of CD138+ plasma cells, producing all classes and subclasses of immunoglobulins with the exception of IgD. These antibody-secreting cells (ASC) are comprised within a larger subset of B cells lacking expression of the complement receptors CD21 and CD35 and sharing the expression of signature genes defining mouse B1 B cells. Single-cell transcriptomic analyses supported the intrathymic differentiation of CD138+ plasma cells alongside other B cell subsets with distinctive molecular phenotypes. Neonatal thymic plasma cells also included clones reactive to pathogenic bacteria that commonly infect children born with antibody deficiency. Thus, our findings point to the thymus as a source of innate humoral immunity in human neonates.

General Microbiology

Immunology

thymus

CD138+ plasma cells

human neonates

The thymus is primarily responsible of the development, maturation and selection of T cells. As recognized more than 30 years ago, this lymphoid organ also houses a functional population of CD19⁺ B cells ¹. In both mice and humans, B cells account for ~ 0.5% of the thymic cellularity ^1,2. While still under investigation, thymic B cells are now known to play an important role in the induction of the central tolerance through the negative selection of self-reactive T cells ^2–5. This function is in accordance with their location in the thymic medulla and the expression of key co-stimulatory molecules such as CD80, CD86 as well as high levels of MHC class II molecules ^2,3,5,6. The autoimmune regulator (AIRE) expressed in mTECs and involved in the negative selective of peripheral antigen-specific T cells was also reported in murine thymic B cells ⁵, supporting a similar role as antigen presenting cells. Additionally, murine thymic B cells were involved in the differentiation of thymocytes into FOXP3-expressing regulatory T cells (Tregs), which also takes place in the medulla ^7,8. Lastly, the presence of class-swtiched antibody-producing thymic B cells was also reported and involved in mechanisms of central tolerance ⁹. Collectively, these animal studies have begun uncovering the important role of thymic B cells in immune regulatory mechanisms. In humans, our understanding of these cells is far less advanced. The disctinction between the two species is particularly relevant because human and mouse thymuses differ structurally ^10,11. Moreover, the aging process of the thymus in mice maintained in conventional laboratory conditions does not reflect physiological conditions ¹⁰.

We recently reported a longitudinal study of the distribution of B cells in human thymuses using specimens from donors aged 5 days to 71 years. Our investigation showed that while B cells were mostly distributed in the thymic medulla in newborns and infants, there was also a progressive accumulation of these cells in the perivascular space (PVS), starting during the first year of life ⁶. The PVS constitutes a third thymic compartment that expands with age concomitantly with the involution of the cortex ^12,13. Furthermore, our studies demonstrated that B cells accumulating with age in the thymic PVS included antibody-secreting cells (ASC) specific to common viruses and vaccine antigens ⁶. The specificity of these ASC to immunizing antigens as well as the kinetics of their accumulation strongly suggested that they resulted from peripheral immune response. These findings revealed an unrecognized role of the thymic PVS as a functional lifetime niche for viral-specific plasma cells (PCs). In the present study, we examined whether functional ASC are already present in the thymus of human newborns presumably before any external antigen exposure. We also investigated their contribution to humoral innate immunity.

The human newborn thymus houses antibody-secreting cells comprised within the CD21 ⁻ CD35 ⁻ B cell subset.

In previous studies, we showed that the human thymus provides a niche supporting antibody-producing plasma cells ⁶. While the number of thymic ASC increases with age, starting during the first year of life, their presence at birth had not been fully evaluated. Here, we characterized ASCs in thymic specimens (n = 12) obtained up to 7 days after birth. This restricted time window ensured that we examined ASCs that had been previously differentiated in utero. Cells secreting IgG ex-vivo without stimulation represented approximately 1 in 10⁴ thymocytes and this frequency was consistent within the first week of life (Fig. 1a, Supplementary Fig. 1a). The spontaneous secretion of immunoglobulins without the need for in vitro stimulation is a hallmark of terminally differentiated plasma cells such as long-lived plasma cells (LLPC) residing in the bone marrow niche. However, while bone marrow LLPC lose the expression of CD19 ¹⁴, thymic ASC were almost exclusively contained in the CD19⁺ fraction (Fig. 1b, Supplementary Fig. 1b). The overall frequency of thymic ASC was equivalent to that of CD19⁺ ASC in adult blood. In contrast, no circulating ASC were detected in cord blood (Fig. 1c, Supplementary Fig. 1c), suggesting that thymic PC developed in situ and did not recirculate during perinatal stages.

The differential expression of CD21 by peripheral blood B cells has been reported to define functionally distinct B cell subsets following influenzae vaccination ¹⁵. Thymic B cells are heterogeneous, with an important subset previously reported to lack expression of both complement receptors CD21 and CD35 ¹. Using these two markers, thymic CD19⁺ B cells could be subdivided into CD21⁻CD35⁻ and CD21⁺CD35⁺ populations (Fig. 1d). CD21⁻CD35⁻ B cells were not present in adult or cord blood. Both subsets were consistently detected in thymus specimens collected within the first 6 months after birth (n = 21; Fig. 1e). Based on these findings, we selected 5 representative thymus specimens ranging in age between 1 day and 4 months for in-depth analysis. All ASC were included in the CD21⁻CD35⁻ subset and spontaneously secreted IgG, IgM, IgA and in some cases IgE (3 out of 5) (Fig. 1f, Supplementary Fig. 1d). While we had previously reported on the presence of IgG, IgM and IgA-secreting cells in the human thymus ⁶, IgE-secreting cells had not been detected. The diversity of immunoglobulin classes produced by neonatal thymic ASC points to active class-switch recombination in utero.

Early studies using transmission electron microscopy (TEM) and immunohistochemistry described subsets of large “asteroid in shape” B cells together with smaller round B cells in the fetal and adult thymic medulla, highlighting the heterogeneity of this population ^16,17. We also used transmission electron microscopy to examine the cellular structure of CD21⁻CD35⁻ and CD21⁺CD35⁺ B cells. Overall, CD21⁻CD35⁻ B cells were morphologically different from CD21⁺CD35⁺ B cells, displaying intracellular organelles and size consistent with activated B cell blasts as well as typical plasma cells with enlarged rough endoplasmic reticulum and cart-wheel shaped nuclear patterns (Fig. 1g, h, Supplementary Fig. 1e). This subset of blastic B cells likely corresponds to the asteroid B cells mentioned previously.

Neonatal thymic CD21 ⁻ CD35 ⁻ and CD21⁺CD35⁺ B cells display a unique transcriptome profile with expression of signature innate-like B cell genes

We next used RNAseq to further characterize neonatal thymic B cell subsets and cord blood B cells as a comparison (Supplementary Table 1, 2). Principal component analysis (PCA) segregated all three populations (Fig. 2a). In addition, unsupervised clustering revealed marked differences between the gene expression signature of thymic CD21⁻CD35⁻ B cells when compared to thymic and cord blood CD21⁺CD35⁺ B cells (Fig. 2b). Among the differentially expressed gene, SPN (sialophorin) encoding CD43 a marker associated with B1 B cells in mice, was upregulated in both thymic B cell subsets when compared to cord blood B cells (Fig. 2c). We then focused on a select set of genes established as the minimal signature of B1 and B2 B cells in mice ^18,19. More than 50% of the genes defining the canonical B1 signature in mice and detected in our transcriptomics analysis, were upregulated in both CD21⁻CD35⁻ and CD21⁺CD35⁺ thymic B cell subsets when compared to cord blood B cells, including ZBTB32, BHLHE41, PLSCR1, GPR55 and MYO1D (Fig. 2d). In contrast, only one gene included in the B2 signature was differentially regulated between these B cell populations (Supplementary Table 3). Of note, we could not detect the expression of AIRE in thymic CD21⁻CD35⁻ or CD21⁺CD35⁺ B cells (Supplementary Tables 1 and 2) as was previously reported for mouse thymic B cells ^2,5,20, suggesting a divergence between the two species.

The neonatal thymic CD21 ⁻ CD35 ⁻ B cell subset is highly heterogenous and includes class-switched cells with a plasma cell phenotype

A total of 1800 genes were differentially expressed between thymic CD21⁺CD35⁺ and CD21⁻CD35⁻ B cell with 1120 upregulated and 680 downregulated genes in the thymic CD21⁻CD35⁻ B cell subset (Supplementary Fig. 2, Supplementary Table 4). Gene set enrichment analysis (GSEA) identified BCR signaling, B cell activation, cell cycle, and defense responses to bacteria among the most upregulated pathways in CD21⁻CD35⁻ B cells (Supplementary Fig. 3a). Several key genes associated with T cell interaction, such as CD80 and CD86, were up-regulated in thymic CD21⁻CD35⁻ B cells (Fig. 3a), supporting a role previously attributed to medullary B cells in T cell negative selection ^2,5. Other activation-induced genes expressed in CD21⁻CD35⁻ B cells included CD30, CD70, CTLA-4 as well as PDCD1, coding for programmed cell death protein 1 (PD1). Moreover, the expression of AICDA further confirmed intrathymic class switch recombination (CSR) previously reported in mice ⁹. Thymic CD21⁻CD35⁻ B cells in neonates also displayed upregulation of genes involved in cell division and proliferation such as MKI67, UBE2C, THYMS, RRM2 and PCNA (Fig. 3a). Moreover, several genes associated with B cell differentiation into plasmablasts and PC were up-regulated in CD21⁻CD35⁻ B cells when compared to CD21⁺CD35⁺ cells. These include CD138, XBP1, MZB1, IL6R, BLIMP1 and BCMA (Fig. 3a). Differential expression of several of these key markers in CD21⁻CD35⁻ B cells was confirmed by high dimensional flow-cytometry (Fig. 3b, Supplementary Fig. 3b). In particular, 50–60% of the CD21⁻CD35⁻ B cells were positive for MKI67, indicating active proliferation of these cells.

In accordance with the upregulation of AICDA and class-switch recombination, the transcriptome profile of CD21⁻CD35⁻ thymic B cells revealed higher expression of genes encoding all classes and subclasses of immunoglubulins with the exception of IGHM and IGHD when compared to CD21⁺CD35⁺ B cells (Fig. 3c). A majority of immunoglobulin heavy (75%), kappa (83%) and lambda (98%) light chains variable region genes were upregulated in CD21⁻CD35⁻ cells with the notable exception of IGHV5.78 that had previously been identified as a pseudogene (Supplementary Fig. 4a). The gene coding for the J chain was also upregulated in the double complement receptor negative B cell subset. Differential membrane expression of these immunoglobulins in CD21⁻CD35⁻ and CD21⁺CD35⁺ cells was verified by flow cytometry (Fig. 3d, Supplementary Fig. 4b). To further confirm these findings, we carried out an in-stage tip proteomics analysis of sorted CD19⁺CD21⁻CD35⁻ and CD19⁺CD21⁺CD35⁺ thymic B cells. The results verified over-expression of IgGA1, IgGA2, IgG1-3 but not IgM in CD21⁻CD35⁻ B cells (Fig. 3e). In addition, the cells that had undergone CSR showed lower levels of IgD.

Intrathymic differentiation of B cells into plasma cells in the thymus of human neonates.

Bulk RNA-sequencing findings suggested a high heterogeneity within CD21⁻CD35⁻ thymic B cell subset. We further characterized these cells using single cell RNA sequencing (scRNA-seq). Data from two donor samples (4-day and 4-month old thymus) were integrated to perform unsupervised community detection based on highly variable genes. The cells were then projected in a two-dimensional space using Uniform Manifold Approximation and Projection (UMAP). This approach revealed 4 different B cell clusters that were comparable between a 4 day-old and a 4 month-old thymus (Fig. 4a, b). The first cluster corresponded to cells expressing genes involved in cell cycle progression and proliferation such as MKI67, UBEC2B, TYMS, HMGB2 and PCNA (Fig. 4c, Supplementary Fig. 5a). These genes matched those reported in Fig. 3a and corresponded to actively dividing cells. The second cluster, smaller in size, included B cells expressing CCL22 and CCL17, two chemokines involved in the chemotaxis of CCR4-expressing Th2 and Tregs ^21,22. B cells in this cluster also expressed the tetraspanin CD9, a marker of IL10-producing regulatory B cells ^23,24 and EBI3, expressed in regulatory plasma cells ²⁵ and involved in CD4 T cell regulation ²⁶ and tolerance ²⁷ (Fig. 4c, Supplementary Fig. 5a). The third and largest cluster was characterized by expression of CD86, CD83, CD72 and CD1C, indicating that B cells in this cluster are well-equipped to interact with T cells ^28–30. This cluster also included B cells expressing markers associated with a mature naïve phenotype, such as IgD ³¹. Lastly, the fourth cluster was characterized by genes expressed in plasmablasts and plasma cells, including SDC1 (CD138), MZB1, XBP1, PRDM1 and JCHAIN (Fig. 4c, Supplementary Fig. 5a). The list of differentially expressed genes for all clusters is provided in Supplementary Table 5.

By analyzing the relative abundance of unspliced and spliced mRNA at the single cell level, we predicted that cluster 2, cluster 3 as well as plasma cell cluster 4 originated from the highly proliferative cluster 1 (Fig. 5a). This cell trajectory analysis using RNA velocity ³² uncovered three destination subsets of thymic B cells, likely endowed with distinct functional properties. The trajectory analysis also revealed two different subgroups of plasma cells in destination cluster 4, expressing either all IGH subclass genes or IGHA and IGHM (Fig. 5a, b). This single-cell RNA velocity analysis provided supportive evidence that B cells differentiate into plasma cells, producing multiple classes and subclasses of immunoglobulins in the thymus of human neonates. These findings concur with the detection of IgM-, IgG, IgA and IgE-ASC within neonatal CD21⁻CD35⁻ thymic B cells as reported in Fig. 1 f.

Thymic CD21^-CD35^-CD138⁺ show evidence of somatic hypermutation (SHM)

The generation of class-switched plasma cells is a central characteristic of B cell responses that typically occur in the germinal centers (GC) in a T cell-dependent manner. Although, CSR can also take place outside of the GCs ^33–35, and without T cell help ^36,37 in what is known as extrafollicular response ³⁸. CSR is governed by activation-induced cytidine deaminase (AID) encoded by the AICDA gene expressed in CD21^-CD35^- thymic B cells. AID is also responsible for the accumulation of SHM in differentiating cells as a means to diversify the antibody repertoire. We hypothesized that CSR of intrathymic B cells could also be accompanied by SHM and possible clonal selection. We conducted a comparative IGHV repertoire analysis of the two major thymic B cells (CD21⁺CD35⁺ and CD21^-CD35^-) as well as the most specific subset of thymic plasma cells CD21^-CD35^-CD138⁺ within the same individuals (n = 5; Supplementary Table 6). First, higher clonality, Simpson clonality, and Simpson’s D indexes ^39,40 were found in CD21^-CD35^- B cells compared to CD35⁺CD35⁺ B cells. Within CD21^-CD35^- B cells, CD138⁺ cells had the highest clonality indexes in all cases (Fig. 6a, Supplementary Fig. 6a, b), suggesting clonal expansion within this cell population. In accordance with these findings, both the iChao1 ⁴¹ and Efron & Thisted ⁴² estimators also indicated a lower diversity within CD138⁺ cells in 5 out the 5 thymus specimens (Supplementary Fig. 6c, d). A pairwise nucleotide sequence comparison using morisita index ⁴³ next revealed some limited level of sequence overlap between thymic CD21^-CD35^- and CD138⁺ cells from the same thymuses but little if any overlap among these populations between the different thymuses (Fig. 6b). The frequency of IGHV rearrangements with somatic mutations appeared higher in thymic CD21^-CD35^-CD138⁺ plasma cells compared to total CD21^-CD35^- or CD35⁺CD35⁺ B cells, although this difference was mostly observed for the 3 month- and 4 month-old thymus specimens, and did not reach statistical significance (Fig. 6c). The average number of SHM per mutated rearrangement was also very low (Fig. 6d). Lastly, the VH usage was similar across the thymuses and groups, with a predominance of rearrangements using IGHV3 and IGHV4 (Supplementary Fig. 6e, f). Together, these results suggested that a restricted contingent of B cells were selected to undergo differentiation into PC intrathymically. While SHM were detected in thymic B cells, these were limited and mostly confined to the CD138⁺ subset.

Neonatal thymic CD138⁺ plasma cells are located in the PVS and secrete immunoglobulins with a reactivity profile characteristic of natural antibodies

Humans are born with a pre-set repertoire of protective natural antibodies (Nabs) assumed to develop without exposure to foreign antigen. A central characteristic of these antibodies is the ability to bind constitutive elements of the bacterial wall, explaining their anti-bacterial properties ^44,45. We hypothesized that thymic ASC differentiating peri-natally could constitute a source of Nabs. To test this hypothesis, we generated recombinant monoclonal antibodies (rAbs) from 362 individual CD21^-CD35^-CD138⁺ isolated from 5 thymus specimens. All rAbs were then tested for their reactivity to bacterial species responsible for recurrent infections in children with agammaglobulinemia such as Staphylococcus aureus and Haemophilus influenzae as well as two additional species Klebsiella pneumoniae, and Escherichia coli. Between 2 to 15% of all PC clones reacted to at least one bacterial species (Fig. 7a, b, Supplementary Fig. 7), with a predominance for reactivity to Gram-positive (S. aureus) over Gram-negative bacteria. We did not detect any clones reactive to K. pneumoniae (Fig. 7a, Supplementary Fig. 7). Albeit limited, this screen underscores the frequency of plasma cells generated in the thymus of neonates and infants that secrete antibodies reactive to bacterial pathogens.

We have previously reported on the propensity of PC to accumulate in the thymic perivascular space (PVS) in infants and adults ⁶. PC generated in neonates also appeared to locate around or in the PVS (Fig. 7c). This specific location may reflect beneficial survival conditions in the PVS for ASC. Moreover, their proximity to vessels supports a contribution of these Nab-producing cells to peripheral immunity in neonates. This particular location may reflect beneficial survival niche conditions for ASC. Moreover, their proximity to vessels supports a contribution of these ASC to peripheral immunity.

In this study we demonstrated that antibody-producing plasma cells are generated in the perinatal human thymus. Functional ASC were observed as early as one day after birth, indicating that the differentiation of these cells started during fetal life. A previous report using human fetal blood at various stages of gestation already provided evidence for CSR and SHM in utero in humans ⁴⁶. Our findings demonstrate that the thymus also supports these fundamental aspects of B cell differentiation.

Our studies also revealed the important heterogeneity of thymic B cells in human neonates, attesting to an intense B cell activity perinatally in this lymphoid organ. We first observed two main populations defined by the expression of the two main complement receptors CD21 and CD35. Based on their small size, structural features seen by TEM and lack of expression of activation markers, CD21⁺CD35⁺ B cells appear as resting cells. In contrast, CD21⁻CD35⁻ B cells are larger, more diverse with respect to their molecular phenotype and express a host of activation and differentiation markers. Remarkably, activated CD21^low B cells had already been reported as a peripheral blood subset of activated B cells prone to further differentiate into plasmablasts and plasma cells following seasonal influenzae vaccination ¹⁵. In neonates, virtually all thymic CD21^−/low cells also lacked expression of CD35, the second complement receptor. Gene expression profiling indicated that the absence of CD21 and CD35 at the cell membrane was due to down-regulation of the corresponding genes, rather than internalization of the receptors. Within the heterogenous CD21⁻CD35⁻ thymic B cell popolution, our scRNAcell analysis revealed 4 distinct subset, the characteristics of which pointed to their possible functions. One subset corresponded to B cells undergoing active proliferation, most likely as the result of stimulation in situ. Based on RNA velocity cell trajectory analysis, this subset of dividing cells led to 3 destination subsets, suggesting that once stimulated, thymic B cells can differentiate into distinct functional phentotypes. The larger destination subset included cells expressing a number of co-stimulatory molecules previously ascribed to cell-to-cell interactions with T cells such as CD80, CD86 and MHC class II molecules. This subset likely includes B cells involved in T cell interaction ^2–5,7. B cells in the second destination subset shared the expression of CD9 with IL-10-producing regulatory B cells ^23,24 as well as the expression of EBI3, a critical subunit of the inhibitory cytokine IL-35 with immunoregulatory plasma cells ²⁵. Moreover, cells within this subset also expressed the CCL22 and CCL17 genes encoding Th2 and Treg-actracting chemokines ^21,22. Specific expression of these two genes in IL-10-secretion Bregs was recently described in the context of antigen-induced arthritis in mice ⁴⁷. It is plausible that differentiated B cells within this subset also display suppressive capabilities and other functional characteristics of Bregs. The last destination cluster included ASCs characterized by a common expression signature of genes involved in differentiation into plasmablasts and plasma cells. Remarkably, the molecular profile of these cells together with the functional characterization of thymic ASC ex vivo, provided evidence of active CSR towards all classes and subclasses of immunoglobulins, including IgE, but excluding IgD. The physiological significance of such broad CSR processes is unclear.

B cells usually undergo CSR and differentiation into plasma cells upon antigen recognition and BCR stimulation. Several converging observations suggest that the differentiation of thymic B cells follow the same mechanism. First, GSEA indentified BCR signaling and B cell activation pathways as up-regulated in thymic ASCs comparatively to other thymic B cells. Second, the higher clonality index observed in CD138⁺ cells compared to other thymic B cell subsets, supports the notion of clonal selection based on the specificity of the BCR. Taken together, these observations suggest that certain thymic B cell clones are selected to undergo differentiation into plasma cells on the basis of their BCR reactivity. As previously reported in mice, these may predominantly include autoreactive clones ^2,5,9.

In mice, B1 B cells are known to develop in the fetal liver and to dominate B cell populations during the first weeks of life ⁴⁸. As a result, most splenic B cells have a B1 cell phenotype in mouse pups ⁴⁹. Through the production of natural antibodies reacting to bacterial carbohydrates, B1 B cells contribution ot protective innate humoral immunity at birth. B1 are well characterized in mice; however, their existence in humans is still questioned. This controversy is primarily due to the lack of a specific set of markers discriminating a human equivalent of mouse B1 B cells. On the other hand, humans are also born with protective natural antibodies, suggesting the existence of a comparable subset of innate-like B cells. Our studies examined whether human neonatal thymic B cells expressed genes included in the canonical B1 and B2 signature recently established in mice ^18,19. Strikingly, 6 out of 10 genes included in the mouse B1 transcriptional signature were upregulated in both thymic B cell subsets when compared to cord blood B cells. In contrast, none of the B2 genes were consistently up-regulated in the same B cell subsets. Thymic B cells also displayed high expression of CD43, a putative marker of B1 B cells ^48,50. While our results do not demonstrate that neonatal thymic B cells correspond to mouse B1 B cells, the similarities in gene expression profiles between the two cell types suggest that they may share certain characteristics. This idea is further supported by the reactivity of immunoglobulin secreted by thymic CD138⁺ plasma cells. Mouse B1 cell-derived antibodies react to pathogen-expressed molecules, including LPS and phosphorylcholine derived from Gram-negative and Gram-positive bacteria respectively ⁵¹. Our studies also revealed that ~ 7% of recombinant monoclonal antibodies generated from CD138⁺ thymic plasma cells reacted to bacteria including Gram-negative bacteria such as H. influenzae and Gram-positive bacteria such as S. aureus, two common pathogenic bacteria infecting children with agammaglobulinemia ⁵².

Nabs present in the serum of non-immunized animals and humans were recognized more than 100 years ago ^53,54. These antibodies are essential elements of natural immunity at birth, providing an innate line of defense against prevalent pathogens ^44,55. Children born with agammaglobulinemia or other forms of antibody deficiency invariably suffer from recurrent bacterial infections unless they receive immunoglobulin replacement therapy ⁵². Despite their importance, the origin of Nabs in human neonates has remained largely unknown. As mentioned above, a significant proportion of neonatal thymic PC also reacted to common pathogenic bacteria. We propose that these cells are an important source of protective Nabs perinatally. Overall, our experiments describe for the first time the intrathymic differentiation of plasma cells secreting natural antibodies in human neonates. The identification of thymic PC clones reactive to pathogenic bacteria commonly infecting children born with primary antibody deficiencies could have important implications for the development of novel replacement therapies.

Sample collection

Thymuses were obtained from neonates and infants undergoing cardiac surgery at the NewYork- Presbyterian Hospital. Cord blood samples were obtained from Carolinas Cord Blood Bank (Duke University). Healthy buffy coats from adults were obtained from the New York Blood Center. This study was approved by the Columbia University institutional review board.

Sample processing

Thymic tissue was collected in cold phosphate-buffered saline (PBS) and washed extensively to remove the blood. A piece of tissue was kept on 10% paraformaldehyde solution for 24h, then placed on 70% ethanol and finally paraffin embedded for histological analysis. The remaining tissue was homogenized using a gentleMACS tissue dissociator (Miltenyi Biotec) and filtered through a 40-µm cell strainer (BD Biosciences). Peripheral blood mononuclear cells (PBMCs) were isolated from cord blood and adult blood by Ficoll density gradient using Ficoll-PaquePLUS (GE HealthCare). Thymocyte and PBMC suspensions were frozen in heat inactivated fetal bovine serum (FBS) containing 10% dimethyl sulfoxide (DMSO, Fisher BioReagents) and kept in liquid nitrogen until use.

Flow cytometry and cell sorting

Cryopreserved cells were thawed into warm RPMI media containing 10% FCS and washed in PBS. B cells were isolated by magnetic cell sorting with EasySep Human B Cell Enrichment Kit (Stem Cell Technologies) following the manufacturer’s instructions and stained in PBS with 2% FCS for 45min with the following fluorochrome conjugated antibodies: anti-CD3 BV786 (Clone SK7, BD Biosciences), anti-CD3 BV570 (Clone UCHT1, Biolegend), anti-CD45 Qdot800 (Clone HI30, Thermo Fisher Scientific), anti-CD19 PECy7 (Clone HIB19, Tonbo Biosciences), anti-CD21 BV711 (B-ly4, BD Biosciences), anti-CD21 PECy5 (B-ly4, BD Biosciences), anti-CD21 V450 (Clone B-ly4, BD Biosciences), anti-CD35 PE (E11, BD Biosciences), anti-CD35 FITC (E11, BD Biosciences), anti-CD38 BV650 (clone HIT2, Biolegend), anti-CD138 PE (clone 44F9, Miltenyi Biotec), anti-CD138 VB515 (clone 44F9, Miltenyi Biotec), anti-CD27 APC Cy7 (clone O323, Tonbo Biosciences), anti-IgG Alexa Fluor 700 (Clone G8-145, BD Biosciences), anti-IgM BV421 (Clone MHM-88, Biolegend), anti-IgA APC (Clone IS11-8E11, Miltenyi Biotec), anti-IgE BV480 (Clone G7-26, BD Biosciences), anti-IgD BV510 (Clone IA6-2, BD Biosciences), anti-CD69 PECy5 (Clone FN50, Biolegend), anti-CD80 BV711 (Clone 2D10, Biolegend), anti-CD86 Alexa Fluor 647 (Clone IT2.2, Biolegend), anti-PD1 PE-Dazzle594 (Clone EH12.2H7, Biolegend), anti-CD39 BV650 (Clone TU66, BD Biosciences), anti-CD59 PE (Clone H19, Biolegend), anti-CD269 PerCPCy5.5 (Clone 19F2, Biolegend), anti-XBP1S PE (Clone Q3-695, BD Biosciences), anti-IRF4 PerCPCy5 (Clone IRF4.3E4, Biolegend), anti-BLIMP1 Alexa Fluor 647 (Clone 6D3, BD Biosciences), anti-Ki67 FITC (Clone SolA15, Thermo Fisher Scientific).

For intracellular staining, cells were fixed and permeabilized using Transcription Factor Staining Buffer Set (eBioscience) following the manufacturer’s instructions prior to staining. Cells were washed in cold PBS with 2% FCS, filtered through a 70µm cell strainer and acquired using BD LSRFortessa or Cytek Aurora flow cytometer. Data were analyzed using FCS Express 6 Research Edition (DeNovo Softaware).

ELISpot assay

To assess the frequency of spontaneous antibody-secreting cells in the thymus of newborns or in cord blood, ELISpot was carried out following the manufacturer’s instructions with some modifications. Briefly, ELISpot plates (MSIPS4510, Millipore-Sigma) were coated with anti-human IgG (7.5µg ml^-1, Mabtech), anti-human IgM (5µg ml^-1, Mabtech), anti-human IgA (5µg ml^-1, Mabtech) and anti-human IgE (5µg ml^-1, Mabtech). After overnight coating and blocking with 1% FCS-PBS for 30min, total thymocytes, PBMCs from cord blood or sorted thymic B cells (CD19⁺CD21⁺CD35⁺ and CD19⁺CD21^-CD35^-) were plated for detection of total ASCs. After overnight incubation at 37^oC, bound antibodies were detected using biotinylated anti-human IgG (1µg ml^-1, Mabtech), anti-human IgM (1µg ml^-1, Mabtech), anti-human IgA (1µg ml^-1, Mabtech) and anti-human IgE (1µg ml^-1, Mabtech). Spots were developed with ELISPOT Blue Color Module (R&D System) using streptavidin-conjugated alkaline phosphatase and 5-bromo-4-chloro-3-indolyl phosphate (BCIP) / nitro blue tetrazolium (NBT) as substrates. Spots were quantified using ELISpot Bioreader 6000 (BIOSYS).

Transmission electron microscopy

CD19⁺CD21⁺CD35⁺ and CD19⁺CD21^-CD35^- cells were sorted on a BDFACS Aria cell sorter and placed in 2% paraformaldehyde/2.5% glutaraldehyde in 0.1M sodium cacodylate fixative buffer, post-fixed with 1% osmium tetroxide followed by 2% uranyl acetate, dehydrated through a graded series of ethanol and embedded in LX112 resin (LADD Research Industries, Burlington, VT). Ultrathin sections were cut using a Leica Ultracut UC7 ultramicrotome (Leica Microsystems), stained with uranyl acetate followed by lead citrate. Grids were examined on a JEOL 1400EX transmission electron microscope at 120 kV. Images were acquired using Gatan Microscopy Suite Software version 2 (Gatan). Images were analyzed using ImageJ software version 1.52a (NIH, USA, https://imagej.nih.gov/ij/).

RNA Sequencing

Thymic B cells (CD19⁺CD21⁺CD35⁺ and CD19⁺CD21^-CD35^-), cord blood and adult blood CD19⁺B cells were sorted as mentioned above and place in lysis buffer (Qiagen). Total RNA extraction was performed RNAeasy Micro Kit (Qiagen) according to the manufacturer’s instructions. RNA QC was done using TapeStation Analysis Software version A.02.02 (Agilent Technologies). Library preparation was performed using the NEBNext Ultra RNA Library Preparation kit with PolyA selection workflow. Libraries were sequenced on a Illumina HiSeq 4000 using a 2x150bp Paired End lengths. Raw sequence data generated was converted into fastq files and de-multiplexed using Illumina's bcl2fastq version 2.17. Raw reads QC was performed using FASTQC. Reads were aligned using STAR aligner v2.5.2b to map the reads to the reference human genome. RNA sequencing data analysis was performed using R Studio version 3.6.0 ⁵⁶ with sva package for batch correction ⁵⁷ and DESEeq2 package for differential expression analysis ⁵⁸. Plots were generated using ggplot2 package ⁵⁹. Gene set enrichment pathways analysis was performed using GSEA software ^60,61.

Proteomics

Sorted CD19⁺CD21⁺CD35⁺ and CD19⁺CD21^-CD35^- cells were analyzed using in-StageTip (iST) method according to the manufacturer’s instructions ⁶². Peptides were lysed, digested, and submitted to MS/MS. The peptides were identified and analyzed using MaxQuant software package (available at http://maxquant.net/maxquant/) and the Perseus software platform was used for statistical analysis. Results are expressed as label-free quantification (LFQ) intensity. Statistical differences were assigned when p<0.05.

Single cell RNA sequencing

Single cell RNA sequencing of sorted CD21^-CD35^-CD19⁺thymic B cells was conducted using Chromium Single Cell 3′ Reagent Kits v2 (10X Genomics) according to the manufacturer’s instructions. The libraries were quantified using KAPA hgDNA Quantification and QC Kit (Kapa Biosystems) and sequenced via NovaSeq 6000 (Illumina). Following the sequencing, the raw data from each sample were demultiplexed, aligned to the GRCh38-1.2.0. human reference genome, and UMI counts were quantified using the 10X Genomics Cell Ranger pipeline (v2.1.1, 10X Genomics). Data analysis was then continued with the filtered barcode matrix files using the Seurat package version 3.0 ⁶³ in R version 3.6.0 ⁵⁶. For the initial QC step, we filtered out the cells that expressed < 200 genes or > 4000 genes for both the donor samples, and any cell that expressed > 4% mitochondrial transcripts content, giving us 2,410 cells from donor 1 (4 days old) and 2,494 cells from donor 2 (4 months old). Gene expression values for each cell were log normalized and scaled by a factor of 10,000. The most variable genes for both donor samples were identified using the FindVariableFeatures function of the Seurat package, which were then used to integrate the two samples as described elsewhere ⁶³. The integrated dataset with 4,904 cells was scaled, centered and used to run PCA analysis of the combined dataset. Based on the PCElbowPlot, we picked a certain number of principal components (PCs) for the clustering analysis when that number reached to the baseline of the standard deviation of PC. The cells were clustered into sub-populations using Seurat’s implementation of a shared nearest neighbor modularity optimization-based clustering algorithm (Louvain’s original algorithm). Cell clusters were visualized using UMAP ⁶⁴. For differential gene expression, we used model-based analysis of single-cell Transcriptomics (MAST) test ⁶⁵ (log fc ≥0.25) and only selected the genes with adjusting p-value <0.05 were used for further GSEA analysis (as described above). RNA velocity analysis was performed using Velocyto pipeline ³² and integrated into the Seurat analysis.

BCR Sequencing

CD21⁺CD35⁺, CD21^-CD35^- and CD21^-CD35^-CD138⁺ cell populations were sorted as described above. DNA extraction was performed using DNeasy Blood and Tissue Kit (Qiagen) following the manufacturer’s instructions and sample was eluted in TE buffer. IGHV sequencing was performed by Adaptive Biotechnologies using the ImmunoSEQ survey level assay. The assay uses 86 primers for the IGHV gene segment, 15 primers for the IGHD gene segment and 7 primers for the IGHJ gene segment ⁶⁶. This generated a fragment capable of identifying the entire spectrum of unique VDJ combinations including functional genes, pseudogenes, and open reading frames. Next, amplicons were sequenced using the Illumina HiSeq platform. The resulting 130 bp sequences permitted inference of the corresponding germline sequences ⁶⁶, and are denoted IGH–VDJ transcripts. A suite of custom algorithms has been developed by Adaptive Biotechnologies to verify, collapse, align and catalog the CDR3 sequences. To assess and remove PCR bias from the multiplex PCR assay, a synthetic immune system with all possible V–J combinations was precisely quantitated as described elsewhere ⁶⁷. The data were subsequently analyzed and visualized using the ImmunoSEQ analyzer provide by Adaptive Biotechnologies.

Recombinant antibody cloning and expression

CD19⁺CD21⁺CD35⁺, CD19⁺CD21^-CD35^- and CD19⁺CD21^-CD35^-CD138⁺ thymic cells were sorted as previously mentioned into 384-well plates containing hypotonic lysis buffer ^68,69 containing 10nM TRIS and 0.75units ml^-1 of RNASin plus (Qiagen) at 4^oC and stored at -80^oC. Next, cDNA was generated using the High-capacity cDNA generation kit (Applied Biosystems) following the manufacturer’s instructions. Multiplexed PCR was performed using primers specific for IGHV, IGKV, IGHC, IGKC, and IGLC gene segments. Resulting amplicons were used as templates for semi-nested PCR to isolate the heavy and light chain genes and to incorporate modifications to allow ligation independent cloning into expression vectors ⁷⁰. Resulting amplicons were inserted into mammalian expression plasmids containing the IGG1 IGKC, or IGLC gene sequence. Recombinant antibodies were then generated by co-transfection of plasmids encoding Ig heavy and light chain pairs into 293FS cells (Invitrogen) using standard polyethylenimine transfection methods ⁷¹.

Quantitation of Ig clone supernatants

The concentration of IgG in the supernatant of 293 cells transfected with plasmids encoding Ig heavy and light chain pairs was assessed by ELISA using the Human IgG ELISA Quantitation kit (E80-104, Bethyl Laboratories) following the manufacturer’s instructions. Optical density was read at 450nm using BioTek Synergy H1 plate reader. Concentration was calculated based on the standard curve for human reference serum and expressed in ng per ml.

Bacterial culture

Staphylococcus aureus GP22, Escherichia coliform (ATCC 25922), Klebsiella pneumoniae KP35⁷², were picked from plated colonies and grown in an overnight culture of 2mL TSB at 37°C. Haemophilus influenzae (ATCC 19418) was grown in a lawn on chocolate plates overnight at 37°C. Bacterial cultures were sub-cultured or re-suspended in TSB until the OD was approximately 0.35 and 10mL of these suspensions were centrifuged at 4,000 x g at 4°C. Pellets were re-suspended in TSB at 10⁹ CFU ml^-1.

Bacterial reactivity of IgG clone supernatants

IgG clone supernatants were assessed for reactivity to four relevant bacteria (see above). Briefly, each IgG clone supernatant was incubated with each bacterial culture for 30min at 37^oC in 96 well cell culture plates (Corning Incorporated). After washing, bacteria were incubated with FITC-conjugated anti-human IgG (Fisher Thermo Scientific) for 30min at RT. After washing, bacteria were fixed in 10% formalin and acquired on a BD LSRFortessa flow cytometer with high-throughput sampling capabilities. Flow cytometry file data were analyzed as described elsewhere in this paper.

Immunofluorescence of paraffin-embedded thymus sections

Tissue sections were deparaffinized in xylene for 10 min, washed with 100% ethanol followed by 95%, 80%, 70% and 50% ethanol, and then rinsed in distilled water. Samples were processed for antigen retrieval, blocking and staining following the Opal Multiplex IHC protocol (PerkinElmer) as described elsewhere⁷³. Anti-CD19 (clone BT51E, NCL-L-CD19-163, Leica Biosystems), anti-CD31 (clone C31.3 + JC/70A, ab199012, Abcam), anti-cytokeratin (clone PCK-26, ab6401, Abcam) and anti-CD138 (clone MI15, PA0088, Leica Biosystems) were titrated and used as primary antibodies. Finally, after DAPI staining, slides were mounted with VECTASHIELD antifade mounting media (Vector Laboratories). Images were taken using Vectra 3.0 Automated Quantitative Pathology Imaging System (PerkinElmer) and inForm® cell analysis software (PerkinElmer). Images were evaluated and validated by an experienced pathologist.

Statistical analysis

The results are presented as mean ± SD and/or normalized z-score unless otherwise specified in the figure legends. Population size is described in the figure legend. All the statistical analyses were performed using GraphPad Prism software version 7.0. Differences were considered statistically significant when p<0.05 using paired/unpair T-test. For RNA Seq, differences were considered statistically significant when Benjamini-Hochberg adjusted p-value > 0.05 and absolute log₂ fold change > 1. Results of statistical tests are listed in Supplementary Tables 1, 2, 3, 4, and 5.

Data availability

The processed data and transcriptome datasets for both RNA-Seq and Single-Cell RNA-Seq generated during this study are available on NCBI GEO with the accession number: GSE152453 and GSE153117, respectively. DNA sequencing data used for BCR repertoire analysis can be accessed at Adaptive Biotechnology ImmunoSEQ Analyzer.

Code availability

Customized scripts generated for this study are available from the corresponding author on request.

Author contribution: H.C. and E.Z. designed and conceived the project. H.C. collected the samples, performed ELISpot, sorting, FC, RNA-Seq, scRNA-Seq, bioinformatics, proteomics, TEM, IF and statistical analysis. R.G.K. and J.F.K. performed single cell cloning. P.D. performed bioinformatics analysis of both RNA-Seq and scRNA-Seq data. A.M.C and A.C.U optimized and quantified the bacteria culture. H.C., C.D., and S.B.S. performed reactivity experiments. S.H. significantly helped to design and analyze FC experiments. E.A.B. and. D.M.K. provided human specimens. H.C. and E.Z. wrote the manuscript. All the authors have read and approved the final manuscript.

Competing of interest: The authors declare no competing interests.

Acknowledgements: We are indebted to Dr. Patricia Morcillo for her help with the TEM study, Dr. Xinzheng Guo for the IF study as well as R. Shihab for his technical assistance and E. Bush as director at the JP Sulzberger Columbia Genome Center. We also thank the Carolinas Cord Blood Bank at Duke University for providing cord blood samples. We are grateful to Drs. Megan Sykes, Donna Farber, Gilles Benichou and Christian Leguern for critical reading of the manuscript.

Funding: This work was funded by NIAID Grant (U01-131339). Flow Cytometry research reported in this publication was performed in the CCTI Flow Cytometry Core, supported in part by the Office of the Director, National Institutes of Health under the award S10OD020056. Single-cell sequencing analysis was funded in part through the NIH/NCI Cancer Center Support Grant P30CA013696. Electron microscopy research were performed at Albert Einstein College of Medicine using JEOL 1400Plus TEM supported through a shared instrumentation Grant (1S10OD016214-01A1). Single-cell cloning was partially funded by NIAID Grant (U01-AI100005). P.D. was supported by a Cancer Research Institute (CRI) Irvington Postdoctoral Fellowship.

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SupplementaryFiguresandCaptions.pdf
Supplementary Figures with Captions
SupplementaryTable1.CordBloodCD19vsThymicCD21CD35.xlsx
Supplementary Table 1
SupplementaryTable2.CordBloodCD19vsThymicCD21CD35.xlsx
Supplementary Table 2
SupplementaryTable3B1signature.xlsx
Supplementary Table 3
SupplementaryTable4.ThymicBcellsubsets.xlsx
Supplementary Table 4
SupplementaryTable5.ListofDEGbycluster.xlsx
Supplementary Table 5
SupplementaryTable6.Numberofproductivetemplates.xlsx
Supplementary Table 6

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Intrathymic differentiation of natural antibody-producing plasma cells in human neonates

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