Acquired nonpermissive BM microenvironment impairs HSC proliferation and maintenance, and B-cell development after HSCT

Melanie de Gier; Jakov Korzhenevich; Franziska M Schmidt; Iga Janowska; Monique M van Ostaijen-ten Dam; Ingrid Pico-Knijnenburg; Davy Cats; Franziska Keller; Petra Hadlova; Mirjam Kunze; Filiz Markfeld-Erol; Kilian Reising; Hailiang Mei; Robbert G M Bredius; Arjan C Lankester; Mirjam van der Burg; Marta Rizzi

doi:10.1182/bloodadvances.2025015915

. 2026 Jan 20;10(8):2786–2800. doi: 10.1182/bloodadvances.2025015915

Acquired nonpermissive BM microenvironment impairs HSC proliferation and maintenance, and B-cell development after HSCT^∗^†

Melanie de Gier ¹, Jakov Korzhenevich ², Franziska M Schmidt ², Iga Janowska ³, Monique M van Ostaijen-ten Dam ¹, Ingrid Pico-Knijnenburg ¹, Davy Cats ⁴, Franziska Keller ², Petra Hadlova ⁵, Mirjam Kunze ⁶, Filiz Markfeld-Erol ⁶, Kilian Reising ⁷, Hailiang Mei ⁴, Robbert G M Bredius ⁸, Arjan C Lankester ⁸, Mirjam van der Burg ^1,^∗, Marta Rizzi ^2,^3,^9,^10,^∗∗

PMCID: PMC13101697 PMID: 41544216

Key Points

•
Nonpermissive BM stroma hinders B-cell recovery after HSCT and rituximab therapy in 1 of 2 twins with X-linked lymphoproliferative disease.
•
Acquired CXCL12 insufficiency in the BM is associated with a pre-BI developmental block, disrupting B-cell differentiation.

Visual Abstract

graphic file with name BLOODA_ADV-2025-015915-ga1.jpg

Abstract

Defects in B-cell reconstitution upon hematopoietic stem cell (HSC) transplant (HSCT) are a common observation, yet the mechanism remains unexplained. The bone marrow (BM) stroma, including mesenchymal stromal cells (MSCs), guides HSC maintenance and B-lymphopoiesis by secreting crucial cytokines. We report acquired, permanent, selective, and complete B-cell deficiency in the context of full donor chimerism in a patient with X-linked lymphoproliferative disease, and aimed to identify the contribution of the BM microenvironment in disrupted B-cell reconstitution after HSCT. We studied longitudinal BM samples from the patient and his identical twin, both of whom underwent HSCT with the same donor with opposite outcomes in B-cell reconstitution. In the BM of the index patient, we observed progressive loss of proliferation of HSCs and a selective block at the pre-BI cell stage. In vitro modeling studies showed limited survival of patient HSCs and a relative accumulation of pre-B cells. Patient-derived MSCs failed to support survival and proliferation of HSCs and B-cell development of healthy HSCs, which was correlated with reduced CXCL12 levels. Using bulk RNA-sequencing of MSCs and in vitro functional studies, we showed global changes in the patients’ MSCs, and a progressive loss of CXCL12 expression. Indeed, survival of the patients’ HSCs improved upon supplementing the in vitro development culture with CXCL12, suggesting a contribution of defective CXCL12 signaling to the phenotype. In summary, our data show that an acquired defect in the BM-stromal microenvironment and exhaustion of HSCs and committed progenitors may cause a permanent nonpermissive state for normal B-cell development.

Introduction

In bone marrow (BM), hematopoietic stem cells (HSCs) give rise to all hematopoietic lineages. The BM stroma cells, like mesenchymal stromal cells (MSCs) and perivascular endothelial cells, support HSCs and determine lineage commitment.¹^,² Because of the substantial overlap between cytokines instructing myeloid and lymphoid specification, their timing and localization is crucial to guide development.³^,⁴ In fact, disturbed BM cytokine milieu results in impaired HSC differentiation.⁵^,⁶

Development of B-cell precursors (BCPs) from HSCs is guided by stage-specific transcription factors and characterized by expression of surface molecules. Specification into the lymphoid compartment is marked by CD62L and CD10 expression, regulated by transcription factors PU.1 and Ikaros.7, 8, 9 CXCR4-mediated positioning near CXCL12 (or stromal cell–derived factor 1)-producing MSCs allows HSCs and later common lymphoid progenitors (CLPs) to receive additional cues like stem cell factor (SCF), Flt-3 ligand (FLT3L), and interleukin-7 (IL-7) to promote B-lymphoid priming.⁵^,¹⁰^,¹¹ IL-7 signaling promotes paired box gene 5 (PAX5), enabling commitment to the progenitor (pro)–B-cell fate, and inducing terminal deoxynucleotidyl (TdT) and recombination activating gene (RAG) expression to initiate immunoglobulin (Ig) heavy chain recombination.12, 13, 14 CD10⁺CD19⁺ pre-BI cells upon expressing Igμ become pre-BII cells. Here, pre–B-cell receptor signaling allows allelic exclusion and light-chain rearrangement. Finally, immature B cells expressing surface IgM egress out of the BM as IgM- and IgD-positive transitional B cells.

Following HSC transplantation (HSCT), B cells emerge from the BM typically after 2 to 3 months; the naive compartment recovers first, whereas the memory B cells require 6 to 24 months.¹⁵^,¹⁶ During this time, defects in B-cell function predispose patients who have undergone HSCT to lymphoproliferative disorder and autoimmune manifestations¹⁷^,¹⁸ that can be treated with B-lymphodepleting agents, for example rituximab (RTX). Although effective, in certain cases B-lymphodepletion persists, and is associated with hypogammaglobulinemia and susceptibility to infections.19, 20, 21, 22, 23

Here, we describe identical twins (patient 1 and patient 2) with X-linked lymphoproliferative disease (XLP), who both received RTX and haplo-identical HSCT from the same donor with opposite outcomes in B-cell reconstitution. Patient 1 failed to achieve complete B-cell reconstitution upon B-cell depletion, whereas patient 2 showed a normal B-cell compartment after HSCT. XLP type 1 is caused by a mutation in the SH2D1A gene (signaling lymphocyte activation molecule-associated protein [SAP]), associated with Epstein-Barr virus (EBV) susceptibility without an intrinsic B-cell defect.²⁴^,²⁵ Four years after transplant, patient 1 had no peripheral B cells and an acquired block at the pre-B stage in the BM, despite normal B-cell reconstitution in previous years. In vitro modeling of B-lymphopoiesis showed that HSCs from patient 1 held the ability to differentiate to immature B cells, but accumulated at the pre-B stage and were unable to sustain B-cell development until the end of the culture. HSC maintenance and B-cell development were similarly impaired when healthy donor (HD) HSCs were cocultured with MSCs from patient 1, pointing to an acquired defect of the BM microenvironment in supporting HSCs and B-lymphopoiesis. In summary, these studies revealed that an acquired nonpermissive stromal environment impairs HSC survival and impedes B-cell development in BM in a RTX-treated post-HSCT setting.

Methods

Patients and controls

BM aspirates from patients with XLP and 6 HD (taken from Vissers et al²⁶) were collected during routine follow-up upon written informed consent as part of biobanking in the Leiden University Medical Center - Willem Alexander Children's Hospital (LUMC-WAKZ) (LUMC ethics committee approval B17.001; supplemental Table 1). CB and BM samples for in vitro B-cell differentiation were obtained in accordance with approval by the Ethics Committee of the University Medical Center Freiburg (353/07_120590 or 20-1109, respectively). BM donors underwent hip replacement surgery and were otherwise healthy.

Flow cytometry of BM

Flow cytometry of BM cells was performed with a 36-color panel previously described (supplemental Table 2).²⁶ Briefly, cryopreserved BM mononuclear cells were thawed and stained with live/dead viability dye FVDUV455, and subsequently with fluorochrome-labeled antibodies (supplemental Methods).

Human progenitor isolation and MSC expansion

CD34⁺ cells from CB and BM were isolated using the CD34 Microbead Kit (Miltenyi Biotec). MSCs were expanded in MesenCult Proliferation medium (Stemcell Technologies) until passage 3 to 5 for subsequent experiments (see the supplemental Methods).

In vitro B-cell differentiation

CD34⁺ cells were expanded for 7 days in complete Iscove modified Dulbecco medium (IMDM; supplemental Methods)²⁷^,²⁸ in U-bottom plates with SCF, FLT3L, and IL-6 (all 25 ng/mL; all Immunotools). On day 7, CD34⁺ cells were kept until day 14 in the presence of SCF, FLT3L, and IL-7, and when indicated CXCL12 (20 ng/mL; Immunotools), or replated on top of 20-Gy irradiated MSCs derived from patients and HDs and kept in cytokine-free complete IMDM (20 000:200 CD34:MSCs cells per well). From day 14, cells were kept in cytokine-free complete IMDM. Differentiation was monitored weekly by flow cytometry (Table 1, “In vitro B-cell differentiation”; supplemental Table 2; supplemental Methods). Intracellular staining was performed on days 14 and 28 with the eBioscience Foxp3/transcription factor kit (Invitrogen, catalog no. 00-5523-00; supplemental Table 2; supplemental Methods). All samples were acquired on the 5L Cytek Aurora (Cytek Biosciences Inc) using Spectroflo software with autofluorescence extraction, and further analyzed in FlowJo (version 10.10.0). All conditions and time points included 5 technical replicates. Populations with <10 cells were excluded from analysis.

Table 1.

Flow cytometry population definitions

Population			Phenotype
Conventional gating
Progenitors			CD34⁺ or CD117⁺
	HSC		CD45RA^˗CD10^˗CD133⁺CD90⁺CD38dimCD49f⁺
	MPP		CD45RA^˗CD10^˗CD133⁺
	EMP		CD45RA^˗CD10^˗CD133^-
	MLP		CD45RA⁺CD10⁺CD133⁺
	BCP		CD45RA⁺CD10⁺CD133^˗
		pro-B	CD34⁺CD19^˗TdT⁺
		pre-BI	CD19⁺cyIgM^˗
		pre-BII	CD19⁺cyIgM⁺
		immB	CD19⁺sIgM⁺
	late GMP		CD45RA⁺CD10^˗CD133^˗
	GMP/LMPP		CD45RA⁺CD10^˗CD133⁺
Myeloid			CD33⁺CD15^+/˗
Erythroid			CD45^˗CD71⁺
B cell			CD19⁺
T cell			CD3⁺
NK cell			CD56⁺CD16^˗lo or CD56^˗/dimCD16⁺
flowSOM clusters
	HSCs		CD45RA^˗CD38lo(CD133⁺)
	MPC		CD45RA^˗CD38^lo/int(CD133^˗)CD33^intCD123⁺
	MPP		CD45RA^intCD38⁺
	MEP		CD45RA^˗CD38^lo/int(CD133^˗)CD36⁺CD71⁺CD105⁺
	BCP
		CLP/pre-pro-B	CD117^loCD127^lo TdT⁺CD10^˗
		pro-B	CD10⁺CD117^loCD127⁺CD79a⁺
		(late) pro-B	CD10⁺CD127^lo CD79a⁺PAX5⁺CD19^int
		pre-BI	CD34^intCD19⁺ CD127⁺ PAX5⁺cyIgM^˗
		pre-BII large cycling	CD34^˗CD19⁺CD20^locyIgM⁺IgM^˗Ki-67⁺
		pre-BII large noncycling	CD19⁺CD20^locyIgM⁺IgM^˗Ki-67^˗
		pre-BII small	CD19⁺CD20^intcyIgM⁺IgM^int
		immB	CD19⁺CD20⁺cyIgM⁺IgM⁺
In vitro B-cell differentiation
	progenitors		CD34⁺
	BCP		CD10⁺CD38⁺
		CLP/pro-B	CD33^˗CD10⁺CD38⁺CD19^˗
		CLP	CD33^˗CD10⁺CD38⁺CD19^˗TdT^˗
		pro-B	CD33^˗CD10⁺CD38⁺CD19^˗TdT⁺
		pre-B	CD33^˗CD10⁺CD38⁺CD19⁺
		pre-BI	CD33^˗CD10⁺CD38⁺CD19⁺PAX5⁺cyIgM^˗
		pre-BII large	CD33^˗CD10⁺CD38⁺CD19⁺PAX5⁺cyIgM⁺Ki-67^highcyCD179a^high
		pre-BII small	CD33^˗CD10⁺CD38⁺CD19⁺PAX5⁺cyIgM⁺Ki-67^lowcyCD179a^low
		immB	CD33^˗CD10⁺CD38⁺CD19⁺IgM⁺
	MPC		CD33⁺

Open in a new tab

EMP, erythro-myeloid progenitor; GMP, granulocyte-macrophage progenitor; immB, immature B; LMPP, lymphoid-primed multipotent progenitor; MEP, megakaryocyte-erythroid progenitor; MLP, multi-lymphoid progenitor; MPC, myeloid-primed hematopoietic precursor; MPP, multipotent progenitor; pre-B, precursor-B; pre-pro-B, pre-progenitor B; pro-B, progenitor-B.

Bulk RNA and ATAC-seq

Total RNA was isolated and submitted for RNA sequencing with ribosomal RNA (rRNA) depletion using NovaSeq 6000 (PE150; Illumina). For assay for transposase-accessible chromatin sequencing (ATAC-seq), 20 000 live cells were delivered to the Leiden Genome Technology Center for library preparation with Tn5 transposase and Nextera XT, and sequenced on the NovaSeq6000 (Illumina; supplemental Methods).

Multiplex immunoassay

Supernatants collected from in vitro B-cell differentiation cocultures on days 14, 21, and 28 and 4-day MSC cultivation were subjected to the Human Hematopoietic Stem Cell LEGENDplex immunoassay (BioLegend) according to the manufacturer’s instructions. Concentrations were obtained in picograms per milliliter. For the nonirradiated MSCs, concentrations were corrected for cell numbers.

Flow cytometry of MSCs

Confluent MSCs were washed in fluorescence-activated cell sorter buffer, and incubated with fluorochrome-labeled antibodies (supplemental Table 2) or unstained controls for 30 minutes at room temperature. In separate tubes, MSCs were prepared for intracellular staining by fixing cells in 4% formaldehyde, permeabilization in perm buffer, Fc-blocking, followed by staining for CXCL12 or isotype controls (supplemental Methods). Cells were then stained for goat anti-mouse (GaM) immunoglobulin and CD90 (both 30 minutes, room temperature; supplemental Table 2; supplemental Methods), and washed before acquisition on the 5-Laser Cytek Aurora with Spectroflo software.

Statistics

Statistical analyses were performed in Prism V10 (GraphPad) with multiple testing correction where applicable.

Results

Description of patients

We describe identical twins, who came into our care when patient 1 developed fulminant infectious mononucleosis, and required ganciclovir, cyclosporin A, etoposide, dexamethasone and RTX (375 mg/m² RTX1; Figure 1A; supplemental Table 1). Genetic screening revealed that both patients had a splice-site mutation of the SH2D1A gene (exon 1 c.117C>T; p.Gly39GlyfsX73), and XLP was diagnosed. To prevent EBV infection and/or fulminant infectious mononucleosis, patient 2 also received RTX (375 mg/m²). In the absence of a matched donor, both patients received purified mobilized CD34⁺ HSCs and ex vivo expanded MSCs²⁹ from the haploidentical father (supplemental Figure 1A) after conditioning with busulfan, cyclophosphamide, and antithymocyte globulin (cumulative dose 150 mg/m², 100 mg/kg, 2.5 mg/kg, respectively). Both patients had normal reconstitution by 12 months, with normal lymphocyte (Figure 1B), CD4⁺ T- and CD19⁺ B-cell numbers (Figure 1C-D), although 2 months after transplant, patient 1 received a single dose of RTX (RTX2; supplemental Figure 1B) for EBV viremia (supplemental Figure 1C). In patient 1, 12 months after HSCT, memory B cells remained low (supplemental Figure 1C), serum IgA was absent (Figure 1E), IgG was low, and the patient required persistent immunoglobulin replacement therapy (IGRT; Figure 1F). Patient 2 developed thrombocytopenia because of mixed chimerism (78% donor) at 9 months after HSCT, and received a HSC boost from the father, resulting in normal recovery (supplemental Figure 1D-F). From the first year after HSCT, both patients had full donor chimerism.

Waning B-cell compartment in the first 4 years after HSCT

During years 2 to 4 after HSCT, in patient 1 peripheral B cells and IgG levels progressively decreased (Figure 1D-F), requiring immunoglobulin replacement therapy (IGRT). A lymph node biopsy revealed follicular hypertrophy and increased IgM⁺ plasma cells in the interfollicular space, with no evidence of malignancy or EBV-induced lymphoproliferation. Four years after transplant, patient 1 developed polyarthritis rheumatoid factor- and antinuclear antibody-positive, coinciding with an increase in serum IgM (Figure 1A,G), which was treated with RTX (RTX3). Hereafter, circulating B cells remained undetectable, despite a HSC boost at 10 years after HSCT from his otherwise healthy and sucessfully transplanted twin brother, and who had full paternal donor chimerism (patient 2; Figure 1A,D). Immunohistochemistry of a BM biopsy from patient 1 at T5 revealed few, albeit detectable, PAX5 and CD79a⁺ cells (data not shown). In both patients, natural killer (NK) cells remained in the low-to-normal range (supplemental Figure 1D), with normal CD4⁺ T-cell numbers (Figure 1C) and normal neutrophils (supplemental Figure 1E), indicating that only the B-cell compartment was affected in patient 1.

In vivo BM B-cell development is blocked at the transition from pre-BI to pre-BII

The progressive loss of peripheral B cells in combination with only a few BCPs in the BM biopsy in patient 1 was suggestive of an acquired defect in BM B-lymphopoiesis. Hence, we analyzed longitudinally collected BM aspirates (T1-T5; Figure 1A; supplemental Table 1) and compared them with BM aspirates from patient 2 and 6 HD. For patient 2, a BM aspirate was taken at 10 years after transplant (supplemental Table 1). In the BM, the proportion of B cells was low at T1, after RTX1, abundant after HSCT (T2-T4) compared with HDs and patient 2, yet almost absent after RTX3 at T5 (Figure 2A), whereas the myeloid, T, NK, and erythroid lines were generally well represented (Figure 2A). At T1 to T4, in patient 1, BCPs constituted 55% to 75% of the CD34-compartment, but only 15% at T5 (Figure 2B). Accordingly, the erythroid-myeloid precursors and granulocyte-macrophage and lymphoid primed multipotent precursors were relatively expanded at T5. Next, we subdivided the BCPs into the developmental stages pro-B, pre-BI, pre-BII, and immature B cells. These subsets were all represented at T1 to T4, and in patient 2. At T5 we found a complete block at the transition from pre-BI to pre-BII cell stage (Figure 2C). Of note, at T1 there was a relative accumulation of pre-BI cells, possibly related to the EBV treatment before HSCT. In summary, after HSCT, 2 RTX treatments, and initial successful reconstitution, patient 1 had a low frequency of BCPs and a block in B-lymphopoiesis at the transition from pre-BI to pre-BII. In contrast, patient 2 showed total BM, CD34⁺, and BCP composition comparable to healthy controls.

Figure 2. — **Longitudinal flow cytometric profiling of patient 1 and patient 2 BM aspirates.** Relative population representation of: (A) total BMMC, and (B) CD34⁺ cells and (C) BCP compartments. (D) UMAP visualization of 100 000 CD34⁺ cells or BCPs from patient 1 T1 to T4, patient 2 and H1 to H6, and 17 239 cells for patient 1 T5 (no B cells) from BM aspirates. Unsupervised hierarchical clustering yielded 14 clusters that could be attributed to the CD34⁺ or BCP compartments. On the right, the UMAP from 6 HDs (blue) is overlaid with the BCPs from patient 1 T5. (E) UMAP of 100 000 CD34⁺ cells or BCPs from HD1 to HD6 overlaid with canonical markers used to identify BCP populations of interest. (F) UMAP of 17 239 CD34⁺ cells or BCPs from patient 1 T5 overlaid with canonical markers used to identify BCP populations of interest. (G) Histogram showing normalized expression of cyIgM in 6 HDs overlaid (blue) compared with patient 1 T5. (H) UMAP showing hierarchical clustering of post-HSCT time points from patient 1 and patient 2 compared with a representative HD (HD3). (I) Histogram displaying expression of Ki-67, CD117, CXCR4, and CD34. (J) Median fluorescence intensity (MFI) of Ki-67, CD117, CXCR4, and CD34 expression on HSC/MPP cells of patient 1 at T1, T3, and T5 compared with a HD. BMMC, BM mononuclear cells; FMO, fluorescence minus 1; UMPA, Uniform Manifold Approximation and Projection.

Patient 1 BCP cells express PAX5 and TdT, but fail to rearrange the immunoglobulin heavy chain in vivo

We dissected the B-developmental block of patient 1 using dimensionality-reduction and unsupervised hierarchical clustering. We identified 14 clusters related to the HSC and BCP compartments (Table 1; Figure 2D; supplemental Figure 2). The Uniform Manifold Approximation and Projection (UMAP) overlay of the 6 HDs with patient 1 at T5 (Figure 2E) showed that HSCs and BCPs were present until the pre-BI stage. T5 BCPs expressed TdT, CD10, PAX5, and CD19, indicating that CLP specification and subsequent pro–B-cell commitment and differentiation to pre-BI was not impaired (Figure 2F). Despite normal PAX5 expression, no further B-cell differentiation stages were observed. Pre-BI cells from HDs expressed cytoplasmic IgM (cyIgM) and the proliferation marker Ki-67 (Figure 2F-G). Patient 1 pre-BI cells failed to express cyIgM as well as Ki-67 (Figure 2F-G). At T4, patient 1 had hypogammaglobulinemia, but the BM showed all BCP stages (Figure 2H). However, the most undifferentiated HSCs, defined by the expression of CD133 (not shown; Figure 2D, orange cluster), were reduced, suggesting a progressive loss of multipotent HSCs (Figure 2H). Importantly, the BCP compartment of patient 2 was comparable to healthy controls (Figure 2H). We characterized the HSCs/multipotent progenitors over time in patient 1, and found a progressive loss of c-Kit (CD117) and Ki-67 expression, indicating a reduction in HSC proliferation and reduced CD34 expression (Figure 2I-J; supplemental Figure 3E). In contrast, CXCR4 expression was markedly increased at T5 (Figure 2I-J; supplemental Figure 3E). In summary, patient BCPs failed to produce cyIgM to become pre-BII cells, despite PAX5 and TdT expression, suggesting an inability to complete variable, diversity, and joining [V(D)J]-recombination. In addition, patient 1 HSCs showed a progressive loss of fitness and function, culminating in absent proliferation and high CXCR4 expression at T5.

Patient 1 HSCs maintain the ability to develop into immature B cells in vitro, but show lower survival and accumulation of pre-B cells

We tested the ability of patient 1 HSCs to develop into B cells in B-cell differentiating conditions.²⁷^,²⁸ In this feeder-free model, CD34⁺ cells isolated from BM were cultured first in the presence of SCF, FLT3L and IL-6 for 1 week; then IL-6 was substituted with IL-7 for another week; and finally cells were kept in culture for up to 6 weeks (Figure 3A; supplemental Figure 3A). The culture with BM CD34⁺ cells isolated from patient 1 at T5 showed reduced survival over time compared with healthy BM CD34⁺ cells (Figure 3B). The CD34⁺ cells showed a defect in the specification into BCPs, marked by the expression of CD10 and CD38. The frequency and numbers of BCPs remained low but detectable along the culture (Figure 3C-D). Hence, the HSCs were not able to sustain the continuous production of BCPs. This was not a general developmental defect as CD33⁺ myeloid precursors were present in normal amounts (Figure 3E). Patient 1 did not have any mature B cells at T5. Therefore, we assessed the ability of T5 HSCs to generate any IgM⁺ B cells in vitro. Indeed, within the few detected BCPs, we observed a normal differentiation-dynamics once the pro–B-cell stage was reached, with progressive appearance of pre- and immature B cells, albeit with a relative accumulation of pre-B cells (Figure 3F). The defect in the HSCs was acquired, as CD34⁺ cells of patient 1 at T1, T3, and patient 2 showed normal specification into BCPs in vitro and comparable distribution of CLP/pro-B, pre-B, and immature B cells (supplemental Figure 3B). Looking at distinct stages of pre–B-cell development by intracellular staining (supplemental Figure 3C-D), we observed normal frequencies of pre-BI, large and small pre-BII at days 14 and 28 of culture from HSCs from T1, T3, and T5 of patient 1 (Figure 3G). Supporting cell identity, lineage marker expression was comparable between samples across defined populations (Figure 3H). Therefore, the block at the pre-BI stage observed in ex vivo staining in T5 could be overcome in vitro in B-cell differentiating conditions (Figure 3G), albeit with a low yield of B cells. Furthermore, at T5, when patient 1 had no B cells, the HSCs maintained the ability to develop into immature B cells in vitro. The very low B-cell yield in the culture is likely the result of reduced proliferation of the HSCs, reduced specification into BCPs, and impaired survival of HSCs and BCPs.

Figure 3. — **In vitro differentiation of BCPs from patient 1 HSCs.** (A) Schematic depiction of B-cell differentiation scheme, canonical surface markers, and experimental approach during the 42-day in vitro B-cell differentiation culture. CD34⁺ cells were expanded from days 0 to 7 supplemented with SCF, FLT3L, and IL-6, and in the presence of SCF, FLT3L, and IL-7 from days 7 to 14, and further cultivated in cytokine-free media up until day 42. Surface staining was performed on days 14, 21, 28, 35, and 42 according to the gating strategy in supplemental Figure 3A. Absolute counts quantified at days 14 to 42 of: (B) live cells, and (C) CD10⁺CD38⁺ BCPs as a percentage of live. Absolute counts of: (D) CD10⁺CD38⁺ BCPs, and (E) CD33⁺ myeloid precursors. (F) Relative frequencies of CD19^˗IgM^˗ CLPs/pro-B cells, pre-B cells, and immature B cells as a percentage of CD10⁺CD38⁺ BCPs. A 95% CI based on 11 HDs (blue) is shown behind the mean and standard error of patient 1 replicates. All data are representative of 5 replicates per sample. (G) Absolute counts and relative proportions of B-cell subsets obtained through intracellular staining. (H) MFI of intracellular markers in pro-B and pre-BI cells. For statistical testing, multiple Brown-Forsythe tests were performed with Holm-Šídák correction, with asterisks indicating per–time point significance as ∗P < .05; ∗∗P < .001. CI, confidence interval; FMO, fluorescence minus 1.

Patient 1–derived MSCs do not support HSC survival and B-cell development of healthy cord blood HSCs

Based on these combined results, we suggested an acquired defect of the BM microenvironment in the support of HSCs and B-cell reconstitution. As a proxy for the BM environment, we generated MSC lines from patients and controls, which were used to support the survival and development of cord blood (CB)-derived healthy CD34⁺ cells into BCPs (Figure 4A). CB-derived CD34⁺ cells cultivated on patient 1 MSCs from T5 showed reduced survival and persistence in culture (Figure 4B-C). Patient 1 MSCs were unable to support the survival and maintenance of myeloid precursors (Figure 4D-E) and BCPs (Figure 4F-G), as shown by the progressive reduction of both CD33⁺ and CD10⁺CD38⁺ cells in culture, respectively. HD-derived MSCs supported the survival and development of CB-CD34⁺ cells into immature B cells, with the progressive appearance of BCPs and pre-B cells by days 21 to 28 and immature cells present at days 35 to 42 (Figure 4H). Within the BCPs developing on MSCs of patient 1 at T5, we found similar frequencies of the distinct B-cell stages, suggesting that patient 1 MSCs mainly affected the survival and proliferation of HSCs, and less the differentiation of committed B cells (Figure 4H). We analyzed the development of BCPs in cultures supported by patient 2 MSCs. Although the overall cell survival was not significantly affected (Figure 4B), the HSCs were reduced, albeit less prominently compared with patient 1 at T5. The distribution of CLP/pro-B, pre-B, and immature B cells was comparable to MSC cultures from HD (Figure 4H). To account for batch differences, we performed a second set of experiments and 5 additional MSC-HDs, and MSCs from T1, T3, T5, and patient 2 that confirmed the impaired ability of the patient 1 MSCs from T5, but also T1, to support the persistence of CD34 HSCs and progenitor cells along the culture and normal dynamic of B-cell differentiation in residual cells (supplemental Figure 4), even though it resulted in higher HSC counts. From this, we proposed that MSCs derived from patient 1 at T5 could not effectively support the survival and maintenance of HSCs and BCPs.

Figure 4. — **Differentiation of HD CB-derived BCPs cocultured with BM-derived MSCs from patient 1.** (A) Schematic depiction of HD CB HSCs cocultured with BM-derived MSCs from patient 1 and a HD. CB CD34⁺ were expanded in SCF, FLT3L, and IL-6 for 7 days, and then replated on top of 20-Gy irradiated MSCs from patient 1 or HD. All conditions were kept in cytokine-free complete IMDM until culture day 42, and surface staining was performed on days 14, 21, 28, 35, and 42. Absolute count from day 14 to 42 of: (B) live cells, and (C) HSCs. CD33⁺ myeloid precursors in: (D) absolute counts, and (E) as a percentage of live. (F) Absolute and (G) relative counts of CD10⁺CD38⁺ BCPs. (H) Relative frequencies of CD19^˗IgM^˗ CLPs/pro-B cells, pre-B cells, and immature B cells as a percentage of CD10⁺CD38⁺ BCPs. Data are based on 5 technical replicates, and are presented as mean with standard error. For statistical testing, 2-way analysis of variance was performed with Holm-Šídák correction with asterisks (∗) indicating per–time point significance as ∗P < .05; ∗∗P < .01; ∗∗∗P < .001; ∗∗∗∗P < .0001.

MSC-transcriptome signatures suggest a role of extracellular matrix in B-cell developmental block

To test whether impaired HSC and BCP maintenance resulted from altered MSC function, we performed bulk RNA and ATAC-seq on MSCs (supplemental Figure 5A-C). The MSCs from our cohort (supplemental Table 1) predominantly expressed genes associated with the osteo- and fibro-lineage, and the adipo-lineage to a lesser extent (Figure 5A). Importantly, the pan-MSC markers were amply expressed, although the expression of leptin-receptor appeared reduced in all patient 1 MSCs. Principal component analysis showed that patient 1 MSCs clustered separately from HD and patient 2, with patient 1 at T1 being the most differential from all samples (supplemental Figure 5C). The top 50 differentially expressed genes also separated patient 1 MSCs from the 6 HD and patient 2 (Figure 5B), with a global difference between patient 1 MSCs and HD MSCs, which did not explain the progressive, acquired loss of B cells after HSCT in patient 1 at T3 to T5. Hence, we performed downstream differentially expressed genes analysis on patient 1 MSCs from T3, T4, and T5 (supplemental Figure 5D). Gene ontology analysis revealed significant enrichment of cellular components and molecular functions associated with extracellular matrix constituents among the downregulated genes (supplemental Figure 5E).

Figure 5. — **Transcriptomic signature of patient-derived MSCs compared with HD.** Log₂-normalized counts and differential gene expression statistics were collected from bulk RNA sequencing reads using the dgeAnalysis R-Shiny application using EdgeR with trimmed mean of M values (TMM) normalization. (A) Log₂-normalized expression of pan-MSC markers (pan-mes), and adipo-, fibro-, and osteo-lineage associated genes. (B) Scaled marker expression of the top 50 most variable genes meeting the log₂ (fold change) cut off of 2. Genes are clustered with unsupervised hierarchical clustering. (C) Immunoassay of cytokines and chemokines involved in hematopoietic support on supernatant collected during days 14 to 28 of cocultured healthy CB CD34⁺ cells and MSCs derived from patient 1 T5 (no B cells), HD3 or no MSC control (˗). Obtained concentrations above the lower limit of quantification were retained and log₂-normalized for visualization. (D) Expression of MSC genes associated with B-cell developmental support. Z scores are calculated from the average expression and standard deviation of all samples. (E) Intracellular staining for CXCL12 in CD90⁺ MSCs of 5 HDs (overlaid, blue) and patient 1 at T5 compared with FMO. (F) Quantification of the MFI of intracellular CXCL12 of 5 HDs (blue) and patient 1 at T5. (G) 12-plex immunoassay of hematopoietic support of cytokines and chemokines accumulated in supernatant nonirradiated MSCs from patient 1 grown to confluency for 4 days. Only values above the limit of quantification were retained. Obtained concentrations in picograms per milliliter were corrected for cell numbers. Z scores are calculated based on the mean and standard deviation of log₂-normalized cell number–corrected concentrations. (H) Ratio of live and progenitor cells and BCPs when BM CD34⁺ cells were expanded in the presence of exogenous CXCL12 and cytokines compared with cytokines only.

Reduction in CXCL12 by differential gene expression and immunoassay analysis

Next, we focused on changes in protein expression, in particular on chemo- and cytokines known to be relevant for hematopoietic niche support. We collected supernatants from the CD34:MSC cocultures at days 14, 21, and 28, and performed unbiased analysis of 12 chemo- and cytokines by multiplex immunoassay. We found that patient 1 T5 MSCs produced similar amounts of IL-6 in supernatant throughout culture days 14 to 28 compared with a representative HD, yet we were unable to detect any production of CXCL12 compared with HD control (Figure 5C). The remaining 10 of 12 cytokines measured were below the detection limit. Targeted RNA-seq data reanalysis of genes associated with B-development revealed increased expression of transforming growth factor β1 in T3 to T5, which is known to impair B-lymphopoiesis.³⁰ We also observed reduced CXCL12 expression in patient 1 MSCs from T3 to T5 (false discovery rate [FDR] = 0.24; Figure 5D) that could not be confirmed by quantitative polymerase chain reaction (not shown). IL-6 expression could not be retrieved from our transcriptomic data. However, ATAC-seq analysis showed similar chromatin accessibility at peaks at the CXCL12 and IL-6 gene region (supplemental Figure 5F), indicating that epigenetic regulation of CXCL12 or IL-6 was similar to controls. Hence, reduction of CXCL12 does not seem to be regulated at the transcription level.

To confirm the levels of CXCL12 protein, we characterized our MSC populations by flow cytometry using canonical pan-MSC markers and intracellular CXCL12. Patient 1, patient 2, and HD MSCs all showed similar expression of all pan-MSC markers (supplemental Figure 5G), but reduced expression of CXCL12 in patient 1 at T5 compared with HDs (Figure 5E-F). CXCL12 release in the supernatant was lower in patient 1 than in HD and patient 2 at all time points analyzed, except for T3 (Figure 5G; supplemental Figure 5H). Few other cytokines reached the limit of quantification, such as IL-7, SCF, IL-11, and IL-15, some of which were also reduced at T5, hence potentially contributing to the reduced fitness of HSCs (supplemental Figure 5H). As a parameter for fitness of MSCs, we measured their doubling time that was comparable for all lines, except for MSCs at T1 and T2 having a longer division time (supplemental Figure 5I). The reduced expression of CXCL12 from MSCs at T5 is supported by the observation of enhanced expression of CXCR4 on HSCs at T5 (Figure 2J). In fact, binding of CXCR4 to CXCL12 results in CXCR4 internalization and partial degradation. We tested the ability of HSCs from T5 to respond to CXCL12 signaling, and supplemented the in vitro development culture with CXCL12, which resulted in improved survival of T5 cultured cells (Figure 5H). HSCs showed increased numbers, confirming their ability to respond to CXCL12. Specification into the BCP did not strongly benefit from the addition of CXCL12. Our data suggest that a relative insufficiency of CXCL12 produced by MSCs may impair the survival and maintenance of HSCs.

Discussion

Integral checkpoints in the differentiation of BCPs are controlled by cytokines and chemokines produced by the supporting cells of the BM microenvironment. This study looked at the survival of HSCs, B-lymphopoiesis and function of MSCs in a patient after HSCT and RTX for XLP, who subsequently became permanently B-cell deficient in the context of full donor chimerism. In the BM, we found a defect in the proliferation of HSCs, a reduction of CLP and pro-B cells, and a developmental block at the pre-BI stage. This phenotype was not observed in the patient’s identical twin brother, who received a transplant from the same donor. This acquired, permanent, selective, and complete B-cell deficiency after HSCT appeared to be associated with a “nonpermissive” BM microenvironment with global changes in the MSCs, and relative CXCL12 insufficiency. Although in the BM, precursor B-cell development at T5 was completely blocked at the transition from pre-BI to pre-BII, despite expression of TdT, CD79a, and PAX5, immature B cells could develop in in vitro B-cell differentiation models, pointing toward a role for niche factors. Survival of HSCs and persistent development of BCPs were defective both in a feeder-free culture of CD34⁺ HSCs from the patient, as well as when HD CB-derived CD34⁺ cells were cocultured with MSCs from the patient. Transcriptome and secretome analysis of patient MSCs revealed that CXCL12 production was slightly reduced compared with HD, corroborating studies showing its role in maintaining precursor viability and differentiation.¹⁰^,³¹

Both of the patients in this study had normal initial recovery of B cells and received transplants for XLP type 1 with a mutation in the SAP-encoding SH2D1A gene. SAP is primarily expressed in NK and T cells as a regulator of signaling lymphocyte activation molecule-receptor signal transduction, and when deficient results in absent cytolytic properties against EBV-infected B cells.32, 33, 34, 35 To date, SH2D1A deficiency has never been associated with defects in B-cell development in BM, or with alterations to BM-MSC homeostasis. Because both patients were full donors from 1 year after HSCT, and patient 2 had otherwise normal B-lymphopoiesis, we infer that the post-HSCT acquired B-developmental defect of patient 1 is irrespective of the underlying condition. Although aberrant B-cell reconstitution following HSCT and/or B-lymphodepletion has been observed in other conditions,19, 20, 21^,²³^,²⁷^,³⁶ only a few have performed in-depth BM analyses to identify the level of the B-developmental defect.²²^,²⁷^,³⁶

The inability of the BM microenvironment to support B-lymphopoiesis has been shown also in other conditions, for example in patients with common variable immunodeficiency²⁷^,³⁷ and in patients with autoimmune vasculitis treated with RTX.³⁶ Similarly, patient 1 B-cell development at T5 after HSCT was blocked at the pre-BI stage, but BCPs could develop up to the immature B-cell stage invitro, albeit with low numbers. Also, our data point to a role for the MSC niche. During development, BCPs are dependent on dynamic levels of SCF, FLT3L, IL-6, and IL-7 to promote differentiation and survival. CXCL12 upon CXCR4-binding is phosphorylated, recruits β-arrestin, and is internalized in the early endosome. CXCR4 can then either recirculate on the surface or be degraded. Chronic CXCL12 stimulus results more likely in degradation of CXCR4 and lower expression.³⁸ We found higher CXCR4 expression in T5 HSCs, supporting the potential contribution to the clinical phenotype of reduced or missing ligand in vivo. CXCL12 can provide phosphoinositide 3-kinase (PI3K) signaling in HSCs, but also docking to CXCL12-rich stromal cells that support survival and differentiation of the HSCs. In fact, HSCs and early BCPs mainly require SCF and IL-6, and are near CXCL12-abundant reticular cells, a subtype of MSCs.39, 40, 41 At the pre-B stage, BCPs egress from IL-7–rich areas, yet still present more CXCR4 molecules on their cell surface.⁵^,³¹^,⁴² In mice, distinct subpopulations of CXCL12-abundant reticular cells have been described with distinct roles in supporting the development of HSCs,³^,⁴^,⁴³ but not in humans.⁴⁰^,⁴⁴^,⁴⁵ We found that defects in HSCs survival and reduced specification into BCPs in vitro was associated with reduced CXCL12 secretion. Although higher serum-CXCL12 corresponds with B-cell replenishment after depletion,⁴⁶ paradoxically, enhanced CXCR4 signaling as in warts, hypogammaglobulinemia, infections, and myelokathexis syndrome also results in B-lymphopenia.⁴⁷^,⁴⁸ Therefore, CXCL12 availability must be tightly regulated across the stages of B-lymphopoiesis in BM to coordinate B-cell development.

In our transcriptomic data, we found a globally disturbed RNA profile in patient 1 MSCs compared with HD and patient 1’s identical twin, which mainly concerned reduction in extracellular matrix–associated proteins, and cytokines and chemokines secreted into the extracellular space. The clinical course of patient 1 is marked with repeated viral (re-)infections and re-activation, conditioning, and lymphodepletion. Hence, several factors may have contributed to the change in function of the MSCs in their abilities to support B-cell development. In fact, even though BM MSCs are generally resistant to apoptosis induction,⁴⁹^,⁵⁰ they can still undergo apoptosis and fail to support HSCs in response to alkylating agents like busulfan and cyclophosphamide.⁵¹ In addition, viral infection with cytomegalovirus has also been shown to impact the immunomodulatory capabilities of BM MSCs.⁵² We hypothesize that both conditioning and viral infection contributed to the disturbed BM niche and negatively impacted the fitness of MSCs in patient 1. Further studies are needed to determine if the CXCL12:CXCR4 axis can be disturbed by such a clinical course. In fact, increased CXCL12:CXCR4 signaling after myeloablation is associated with better regeneration of the HSC niche.⁵³^,⁵⁴

In conclusion, we show that aberrant B-cell reconstitution in a patient receiving RTX and HSCT for XLP can be attributed, at least in part, to an acquired defect in CXCL12:CXCR4 signaling that was not seen in the patient’s identical twin brother. Although the clinical course of these patients can be deemed highly patient-specific and nongeneralizable, the understanding of incomplete B-cell or humoral recovery after HSCT and/or B-lymphodepletion is currently limited due to, in principle, good clinical management with IGRT not necessitating further research or acquisition of further BM samples. In-depth BM analyses of such patients can provide us with valuable insights into the delicate rheostat mechanisms that facilitate or impair BM B-cell reconstitution and development in general. Here, to the best of our knowledge, we describe for the first time an acquired defect in HSC maintenance and precursor B-cell development caused by a nonpermissive BM-stromal microenvironment.

Conflict-of-interest disclosure: The authors declare no competing financial interests.

Acknowledgments

M.d.G. has received financial support from the European Society for Immunodeficiencies, European Federation of Immunological Societies-Immunology Letters, and received a Royal Netherlands Academy of Arts and Sciences Ter Meulen grant for this transnational collaboration. M.v.d.B. has received funding from CSL Behring. M.R. was funded by the Deutsche Forschungsgemeinschaft (German Research Foundation, project number 468499998).

Authorship

Contribution: M.d.G., A.C.L., M.v.d.B., and M.R. conceived the project, designed, and interpreted experiments; experiments were performed by M.d.G., with technical assistance from J.K., F.M.S., I.J., M.M.v.O.-t.D., I.P.-K., and F.K.; M.d.G. analyzed the data with support from J.K., F.M.S., I.J., D.C., J.K., H.M., A.C.L, P.H., M.R., and M.v.d.B.; M.K., F.M.-E., K.R., R.G.M.B., and A.C.L. included patients and collected samples; and M.d.G., A.C.L., M.v.d.B., and M.R. wrote the manuscript.

Footnotes

^∗

J.K. and F.M.S. contributed equally to this study.

^†

A.C.L., M.v.d.B., and M.R. are joint senior authors.

Bulk RNA and assay for transposase-accessible chromatin sequencing data have been deposited at https://ega-archive.org/datasets/EGAD50000002072.

Flow cytometry data and further raw data files are available from the corresponding authors, Mirjam van der Burg (m.van_der_burg@lumc.nl) and Marta Rizzi (marta.rizzi@uniklinik-freiburg.de), on request.

The full-text version of this article contains a data supplement.

Contributor Information

Mirjam van der Burg, Email: m.van_der_burg@lumc.nl.

Marta Rizzi, Email: marta.rizzi@uniklinik-freiburg.de.

Supplementary Material

Supplemental Methods, Tables, and Figures

BLOODA_ADV-2025-015915-mmc1.pdf^{(2.7MB, pdf)}

References

1.Morrison SJ, Scadden DT. The bone marrow niche for haematopoietic stem cells. Nature. 2014;505(7483):327–334. doi: 10.1038/nature12984. [DOI] [PMC free article] [PubMed] [Google Scholar]
2.Sugiyama T, Omatsu Y, Nagasawa T. Niches for hematopoietic stem cells and immune cell progenitors. Int Immunol. 2019;31(1):5–11. doi: 10.1093/intimm/dxy058. [DOI] [PubMed] [Google Scholar]
3.Tikhonova AN, Dolgalev I, Hu H, et al. The bone marrow microenvironment at single-cell resolution. Nature. 2019;569(7755):222–228. doi: 10.1038/s41586-019-1104-8. [DOI] [PMC free article] [PubMed] [Google Scholar]
4.Baccin C, Al-Sabah J, Velten L, et al. Combined single-cell and spatial transcriptomics reveal the molecular, cellular and spatial bone marrow niche organization. Nat Cell Biol. 2020;22(1):38–48. doi: 10.1038/s41556-019-0439-6. [DOI] [PMC free article] [PubMed] [Google Scholar]
5.Cordeiro Gomes A, Hara T, Lim VY, et al. Hematopoietic stem cell niches produce lineage-instructive signals to control multipotent progenitor differentiation. Immunity. 2016;45(6):1219–1231. doi: 10.1016/j.immuni.2016.11.004. [DOI] [PMC free article] [PubMed] [Google Scholar]
6.Comazzetto S, Murphy MM, Berto S, Jeffery E, Zhao Z, Morrison SJ. Restricted hematopoietic progenitors and erythropoiesis require SCF from leptin receptor+ niche cells in the bone marrow. Cell Stem Cell. 2019;24(3) doi: 10.1016/j.stem.2018.11.022. [DOI] [PMC free article] [PubMed] [Google Scholar]
7.Thompson RC, Herscovitch M, Zhao I, Ford TJ, Gilmore TD. NF-kappaB down-regulates expression of the B-lymphoma marker CD10 through a miR-155/PU.1 pathway. J Biol Chem. 2011;286(3):1675–1682. doi: 10.1074/jbc.M110.177063. [DOI] [PMC free article] [PubMed] [Google Scholar]
8.Kohn LA, Hao QL, Sasidharan R, et al. Lymphoid priming in human bone marrow begins before expression of CD10 with upregulation of L-selectin. Nat Immunol. 2012;13(10):963–971. doi: 10.1038/ni.2405. [DOI] [PMC free article] [PubMed] [Google Scholar]
9.Pérez-Vera P, Reyes-León A, Fuentes-Pananá EM. Signaling proteins and transcription factors in normal and malignant early B cell development. Bone Marrow Res. 2011;2011 doi: 10.1155/2011/502751. [DOI] [PMC free article] [PubMed] [Google Scholar]
10.Fistonich C, Zehentmeier S, Bednarski JJ, et al. Cell circuits between B cell progenitors and IL-7+ mesenchymal progenitor cells control B cell development. J Exp Med. 2018;215(10):2586–2599. doi: 10.1084/jem.20180778. [DOI] [PMC free article] [PubMed] [Google Scholar]
11.Petkau G, Turner M. Signalling circuits that direct early B-cell development. Biochem J. 2019;476(5):769–778. doi: 10.1042/BCJ20180565. [DOI] [PubMed] [Google Scholar]
12.Kaiser FMP, Janowska I, Menafra R, et al. IL-7 receptor signaling drives human B-cell progenitor differentiation and expansion. Blood. 2023;142(13):1113–1130. doi: 10.1182/blood.2023019721. [DOI] [PMC free article] [PubMed] [Google Scholar]
13.Ramírez J, Lukin K, Hagman J. From hematopoietic progenitors to B cells: mechanisms of lineage restriction and commitment. Curr Opin Immunol. 2010;22(2):177–184. doi: 10.1016/j.coi.2010.02.003. [DOI] [PMC free article] [PubMed] [Google Scholar]
14.Zandi S, Mansson R, Tsapogas P, Zetterblad J, Bryder D, Sigvardsson M. EBF1 is essential for B-lineage priming and establishment of a transcription factor network in common lymphoid progenitors. J Immunol. 2008;181(5):3364–3372. doi: 10.4049/jimmunol.181.5.3364. [DOI] [PubMed] [Google Scholar]
15.van der Maas NG, Berghuis D, van der Burg M, Lankester AC. B cell reconstitution and influencing factors after hematopoietic stem cell transplantation in children. Front Immunol. 2019;10:782. doi: 10.3389/fimmu.2019.00782. [DOI] [PMC free article] [PubMed] [Google Scholar]
16.Zhou G, Zhan Q, Huang L, et al. The dynamics of B-cell reconstitution post allogeneic hematopoietic stem cell transplantation: a real-world study. J Intern Med. 2024;295(5):634–650. doi: 10.1111/joim.13776. [DOI] [PubMed] [Google Scholar]
17.Lum SH, Elfeky R, Achini FR, et al. Outcome of non-hematological autoimmunity after hematopoietic cell transplantation in children with primary immunodeficiency. J Clin Immunol. 2021;41(1):171–184. doi: 10.1007/s10875-020-00895-3. [DOI] [PubMed] [Google Scholar]
18.Su YY, Yu YF, Yan ZY, et al. Post-transplant lymphoproliferative disorders after allogeneic hematopoietic stem cell transplantation: a case report, meta-analysis, and systematic review. Diagn Pathol. 2024;19(1):122. doi: 10.1186/s13000-024-01544-8. [DOI] [PMC free article] [PubMed] [Google Scholar]
19.Ott de Bruin LM, Pico-Knijnenburg I, van Ostaijen-Ten Dam MM, et al. Persistent hypogammaglobulinemia after receiving rituximab post-HSCT is not caused by an intrinsic B cell defect. Int J Mol Sci. 2023;24(21):16012. doi: 10.3390/ijms242116012. [DOI] [PMC free article] [PubMed] [Google Scholar]
20.Marzollo A, Serena T, Mainardi C, et al. Hypogammaglobulinemia in children after hematopoietic stem cell transplantation and rituximab treatment: relevance of B cell subsets. J Pediatr Hematol Oncol. 2023;45(1):e145–e149. doi: 10.1097/MPH.0000000000002582. [DOI] [PubMed] [Google Scholar]
21.Makatsori M, Kiani-Alikhan S, Manson AL, et al. Hypogammaglobulinaemia after rituximab treatment-incidence and outcomes. Qjm. 2014;107(10):821–828. doi: 10.1093/qjmed/hcu094. [DOI] [PubMed] [Google Scholar]
22.Levy R, Mahévas M, Galicier L, et al. Profound symptomatic hypogammaglobulinemia: a rare late complication after rituximab treatment for immune thrombocytopenia. Report of 3 cases and systematic review of the literature. Autoimmun Rev. 2014;13(10):1055–1063. doi: 10.1016/j.autrev.2014.08.036. [DOI] [PubMed] [Google Scholar]
23.Luterbacher F, Bernard F, Baleydier F, Ranza E, Jandus P, Blanchard-Rohner G. Case report: persistent hypogammaglobulinemia more than 10 years after rituximab given post-HSCT. Front Immunol. 2021;12 doi: 10.3389/fimmu.2021.773853. [DOI] [PMC free article] [PubMed] [Google Scholar]
24.Booth C, Gilmour KC, Veys P, et al. X-linked lymphoproliferative disease due to SAP/SH2D1A deficiency: a multicenter study on the manifestations, management and outcome of the disease. Blood. 2011;117(1):53–62. doi: 10.1182/blood-2010-06-284935. [DOI] [PMC free article] [PubMed] [Google Scholar]
25.Pachlopnik Schmid J, Canioni D, Moshous D, et al. Clinical similarities and differences of patients with X-linked lymphoproliferative syndrome type 1 (XLP-1/SAP deficiency) versus type 2 (XLP-2/XIAP deficiency) Blood. 2011;117(5):1522–1529. doi: 10.1182/blood-2010-07-298372. [DOI] [PubMed] [Google Scholar]
26.Vissers LTW, van Ostaijen-Ten Dam MM, Melsen JE, et al. Potential role of B- and NK-cells in the pathogenesis of pediatric aplastic anemia through deep phenotyping. Front Immunol. 2024;15 doi: 10.3389/fimmu.2024.1328175. [DOI] [PMC free article] [PubMed] [Google Scholar]
27.Troilo A, Wehr C, Janowska I, et al. Nonpermissive bone marrow environment impairs early B-cell development in common variable immunodeficiency. Blood. 2020;135(17):1452–1457. doi: 10.1182/blood.2019003855. [DOI] [PMC free article] [PubMed] [Google Scholar]
28.Kraus H, Kaiser S, Aumann K, et al. A feeder-free differentiation system identifies autonomously proliferating B cell precursors in human bone marrow. J Immunol. 2014;192(3):1044–1054. doi: 10.4049/jimmunol.1301815. [DOI] [PubMed] [Google Scholar]
29.Ball LM, Bernardo ME, Roelofs H, et al. Cotransplantation of ex vivo expanded mesenchymal stem cells accelerates lymphocyte recovery and may reduce the risk of graft failure in haploidentical hematopoietic stem-cell transplantation. Blood. 2007;110(7):2764–2767. doi: 10.1182/blood-2007-04-087056. [DOI] [PubMed] [Google Scholar]
30.Lebman DA, Edmiston JS. The role of TGF-beta in growth, differentiation, and maturation of B lymphocytes. Microbes Infect. 1999;1(15):1297–1304. doi: 10.1016/s1286-4579(99)00254-3. [DOI] [PubMed] [Google Scholar]
31.Zehentmeier S, Pereira JP. Cell circuits and niches controlling B cell development. Immunol Rev. 2019;289(1):142–157. doi: 10.1111/imr.12749. [DOI] [PMC free article] [PubMed] [Google Scholar]
32.Dupré L, Andolfi G, Tangye SG, et al. SAP controls the cytolytic activity of CD8+ T cells against EBV-infected cells. Blood. 2005;105(11):4383–4389. doi: 10.1182/blood-2004-08-3269. [DOI] [PubMed] [Google Scholar]
33.Wang N, Satoskar A, Faubion W, et al. The cell surface receptor SLAM controls T cell and macrophage functions. J Exp Med. 2004;199(9):1255–1264. doi: 10.1084/jem.20031835. [DOI] [PMC free article] [PubMed] [Google Scholar]
34.Bottino C, Falco M, Parolini S, et al. NTB-A [correction of GNTB-A], a novel SH2D1A-associated surface molecule contributing to the inability of natural killer cells to kill Epstein-Barr virus-infected B cells in X-linked lymphoproliferative disease. J Exp Med. 2001;194(3):235–246. doi: 10.1084/jem.194.3.235. [DOI] [PMC free article] [PubMed] [Google Scholar]
35.Cannons JL, Tangye SG, Schwartzberg PL. SLAM family receptors and SAP adaptors in immunity. Annu Rev Immunol. 2011;29:665–705. doi: 10.1146/annurev-immunol-030409-101302. [DOI] [PubMed] [Google Scholar]
36.Thiel J, Schmidt FM, Lorenzetti R, et al. Defects in B-lymphopoiesis and B-cell maturation underlie prolonged B-cell depletion in ANCA-associated vasculitis. Ann Rheum Dis. 2024;83(11):1536–1548. doi: 10.1136/ard-2024-225587. [DOI] [PMC free article] [PubMed] [Google Scholar]
37.Ochtrop ML, Goldacker S, May AM, et al. T and B lymphocyte abnormalities in bone marrow biopsies of common variable immunodeficiency. Blood. 2011;118(2):309–318. doi: 10.1182/blood-2010-11-321695. [DOI] [PubMed] [Google Scholar]
38.Marchese A. Endocytic trafficking of chemokine receptors. Curr Opin Cell Biol. 2014;27:72–77. doi: 10.1016/j.ceb.2013.11.011. [DOI] [PMC free article] [PubMed] [Google Scholar]
39.Nakatani T, Sugiyama T, Omatsu Y, Watanabe H, Kondoh G, Nagasawa T. Ebf3+ niche-derived CXCL12 is required for the localization and maintenance of hematopoietic stem cells. Nat Commun. 2023;14(1):6402. doi: 10.1038/s41467-023-42047-2. [DOI] [PMC free article] [PubMed] [Google Scholar]
40.Aoki K, Kurashige M, Ichii M, et al. Identification of CXCL12-abundant reticular cells in human adult bone marrow. Br J Haematol. 2021;193(3):659–668. doi: 10.1111/bjh.17396. [DOI] [PMC free article] [PubMed] [Google Scholar]
41.Gerosa RC, Boettcher S, Kovtonyuk LV, et al. CXCL12-abundant reticular cells are the major source of IL-6 upon LPS stimulation and thereby regulate hematopoiesis. Blood Adv. 2021;5(23):5002–5015. doi: 10.1182/bloodadvances.2021005531. [DOI] [PMC free article] [PubMed] [Google Scholar]
42.Beck TC, Gomes AC, Cyster JG, Pereira JP. CXCR4 and a cell-extrinsic mechanism control immature B lymphocyte egress from bone marrow. J Exp Med. 2014;211(13):2567–2581. doi: 10.1084/jem.20140457. [DOI] [PMC free article] [PubMed] [Google Scholar]
43.Nakashima K, Zhou X, Kunkel G, et al. The novel zinc finger-containing transcription factor osterix is required for osteoblast differentiation and bone formation. Cell. 2002;108(1):17–29. doi: 10.1016/s0092-8674(01)00622-5. [DOI] [PubMed] [Google Scholar]
44.Tormin A, Li O, Brune JC, et al. CD146 expression on primary nonhematopoietic bone marrow stem cells is correlated with in situ localization. Blood. 2011;117(19):5067–5077. doi: 10.1182/blood-2010-08-304287. [DOI] [PMC free article] [PubMed] [Google Scholar]
45.Li H, Ghazanfari R, Zacharaki D, et al. Low/negative expression of PDGFR-α identifies the candidate primary mesenchymal stromal cells in adult human bone marrow. Stem Cell Rep. 2014;3(6):965–974. doi: 10.1016/j.stemcr.2014.09.018. [DOI] [PMC free article] [PubMed] [Google Scholar]
46.Dunleavy K, Hakim F, Kim HK, et al. B-cell recovery following rituximab-based therapy is associated with perturbations in stromal derived factor-1 and granulocyte homeostasis. Blood. 2005;106(3):795–802. doi: 10.1182/blood-2004-08-3198. [DOI] [PMC free article] [PubMed] [Google Scholar]
47.Zehentmeier S, Lim VY, Ma Y, et al. Dysregulated stem cell niches and altered lymphocyte recirculation cause B and T cell lymphopenia in WHIM syndrome. Sci Immunol. 2022;7(75) doi: 10.1126/sciimmunol.abo3170. [DOI] [PMC free article] [PubMed] [Google Scholar]
48.Mc Guire PJ, Cunningham-Rundles C, Ochs H, Diaz GA. Oligoclonality, impaired class switch and B-cell memory responses in WHIM syndrome. Clin Immunol. 2010;135(3):412–421. doi: 10.1016/j.clim.2010.02.006. [DOI] [PMC free article] [PubMed] [Google Scholar]
49.Mueller LP, Luetzkendorf J, Mueller T, Reichelt K, Simon H, Schmoll HJ. Presence of mesenchymal stem cells in human bone marrow after exposure to chemotherapy: evidence of resistance to apoptosis induction. Stem Cells. 2006;24(12):2753–2765. doi: 10.1634/stemcells.2006-0108. [DOI] [PubMed] [Google Scholar]
50.Li J, Law HK, Lau YL, Chan GC. Differential damage and recovery of human mesenchymal stem cells after exposure to chemotherapeutic agents. Br J Haematol. 2004;127(3):326–334. doi: 10.1111/j.1365-2141.2004.05200.x. [DOI] [PubMed] [Google Scholar]
51.Kemp K, Morse R, Sanders K, Hows J, Donaldson C. Alkylating chemotherapeutic agents cyclophosphamide and melphalan cause functional injury to human bone marrow-derived mesenchymal stem cells. Ann Hematol. 2011;90(7):777–789. doi: 10.1007/s00277-010-1141-8. [DOI] [PubMed] [Google Scholar]
52.Meisel R, Heseler K, Nau J, et al. Cytomegalovirus infection impairs immunosuppressive and antimicrobial effector functions of human multipotent mesenchymal stromal cells. Mediators Inflamm. 2014;2014:898630. doi: 10.1155/2014/898630. [DOI] [PMC free article] [PubMed] [Google Scholar]
53.Lévesque JP, Hendy J, Takamatsu Y, Simmons PJ, Bendall LJ. Disruption of the CXCR4/CXCL12 chemotactic interaction during hematopoietic stem cell mobilization induced by GCSF or cyclophosphamide. J Clin Invest. 2003;111(2):187–196. doi: 10.1172/JCI15994. [DOI] [PMC free article] [PubMed] [Google Scholar]
54.Ponomaryov T, Peled A, Petit I, et al. Induction of the chemokine stromal-derived factor-1 following DNA damage improves human stem cell function. J Clin Invest. 2000;106(11):1331–1339. doi: 10.1172/JCI10329. [DOI] [PMC free article] [PubMed] [Google Scholar]

Associated Data

This section collects any data citations, data availability statements, or supplementary materials included in this article.

Supplementary Materials

Supplemental Methods, Tables, and Figures

BLOODA_ADV-2025-015915-mmc1.pdf^{(2.7MB, pdf)}

[bib1] 1.Morrison SJ, Scadden DT. The bone marrow niche for haematopoietic stem cells. Nature. 2014;505(7483):327–334. doi: 10.1038/nature12984. [DOI] [PMC free article] [PubMed] [Google Scholar]

[bib2] 2.Sugiyama T, Omatsu Y, Nagasawa T. Niches for hematopoietic stem cells and immune cell progenitors. Int Immunol. 2019;31(1):5–11. doi: 10.1093/intimm/dxy058. [DOI] [PubMed] [Google Scholar]

[bib3] 3.Tikhonova AN, Dolgalev I, Hu H, et al. The bone marrow microenvironment at single-cell resolution. Nature. 2019;569(7755):222–228. doi: 10.1038/s41586-019-1104-8. [DOI] [PMC free article] [PubMed] [Google Scholar]

[bib4] 4.Baccin C, Al-Sabah J, Velten L, et al. Combined single-cell and spatial transcriptomics reveal the molecular, cellular and spatial bone marrow niche organization. Nat Cell Biol. 2020;22(1):38–48. doi: 10.1038/s41556-019-0439-6. [DOI] [PMC free article] [PubMed] [Google Scholar]

[bib5] 5.Cordeiro Gomes A, Hara T, Lim VY, et al. Hematopoietic stem cell niches produce lineage-instructive signals to control multipotent progenitor differentiation. Immunity. 2016;45(6):1219–1231. doi: 10.1016/j.immuni.2016.11.004. [DOI] [PMC free article] [PubMed] [Google Scholar]

[bib6] 6.Comazzetto S, Murphy MM, Berto S, Jeffery E, Zhao Z, Morrison SJ. Restricted hematopoietic progenitors and erythropoiesis require SCF from leptin receptor+ niche cells in the bone marrow. Cell Stem Cell. 2019;24(3) doi: 10.1016/j.stem.2018.11.022. [DOI] [PMC free article] [PubMed] [Google Scholar]

[bib7] 7.Thompson RC, Herscovitch M, Zhao I, Ford TJ, Gilmore TD. NF-kappaB down-regulates expression of the B-lymphoma marker CD10 through a miR-155/PU.1 pathway. J Biol Chem. 2011;286(3):1675–1682. doi: 10.1074/jbc.M110.177063. [DOI] [PMC free article] [PubMed] [Google Scholar]

[bib8] 8.Kohn LA, Hao QL, Sasidharan R, et al. Lymphoid priming in human bone marrow begins before expression of CD10 with upregulation of L-selectin. Nat Immunol. 2012;13(10):963–971. doi: 10.1038/ni.2405. [DOI] [PMC free article] [PubMed] [Google Scholar]

[bib9] 9.Pérez-Vera P, Reyes-León A, Fuentes-Pananá EM. Signaling proteins and transcription factors in normal and malignant early B cell development. Bone Marrow Res. 2011;2011 doi: 10.1155/2011/502751. [DOI] [PMC free article] [PubMed] [Google Scholar]

[bib10] 10.Fistonich C, Zehentmeier S, Bednarski JJ, et al. Cell circuits between B cell progenitors and IL-7+ mesenchymal progenitor cells control B cell development. J Exp Med. 2018;215(10):2586–2599. doi: 10.1084/jem.20180778. [DOI] [PMC free article] [PubMed] [Google Scholar]

[bib11] 11.Petkau G, Turner M. Signalling circuits that direct early B-cell development. Biochem J. 2019;476(5):769–778. doi: 10.1042/BCJ20180565. [DOI] [PubMed] [Google Scholar]

[bib12] 12.Kaiser FMP, Janowska I, Menafra R, et al. IL-7 receptor signaling drives human B-cell progenitor differentiation and expansion. Blood. 2023;142(13):1113–1130. doi: 10.1182/blood.2023019721. [DOI] [PMC free article] [PubMed] [Google Scholar]

[bib13] 13.Ramírez J, Lukin K, Hagman J. From hematopoietic progenitors to B cells: mechanisms of lineage restriction and commitment. Curr Opin Immunol. 2010;22(2):177–184. doi: 10.1016/j.coi.2010.02.003. [DOI] [PMC free article] [PubMed] [Google Scholar]

[bib14] 14.Zandi S, Mansson R, Tsapogas P, Zetterblad J, Bryder D, Sigvardsson M. EBF1 is essential for B-lineage priming and establishment of a transcription factor network in common lymphoid progenitors. J Immunol. 2008;181(5):3364–3372. doi: 10.4049/jimmunol.181.5.3364. [DOI] [PubMed] [Google Scholar]

[bib15] 15.van der Maas NG, Berghuis D, van der Burg M, Lankester AC. B cell reconstitution and influencing factors after hematopoietic stem cell transplantation in children. Front Immunol. 2019;10:782. doi: 10.3389/fimmu.2019.00782. [DOI] [PMC free article] [PubMed] [Google Scholar]

[bib16] 16.Zhou G, Zhan Q, Huang L, et al. The dynamics of B-cell reconstitution post allogeneic hematopoietic stem cell transplantation: a real-world study. J Intern Med. 2024;295(5):634–650. doi: 10.1111/joim.13776. [DOI] [PubMed] [Google Scholar]

[bib17] 17.Lum SH, Elfeky R, Achini FR, et al. Outcome of non-hematological autoimmunity after hematopoietic cell transplantation in children with primary immunodeficiency. J Clin Immunol. 2021;41(1):171–184. doi: 10.1007/s10875-020-00895-3. [DOI] [PubMed] [Google Scholar]

[bib18] 18.Su YY, Yu YF, Yan ZY, et al. Post-transplant lymphoproliferative disorders after allogeneic hematopoietic stem cell transplantation: a case report, meta-analysis, and systematic review. Diagn Pathol. 2024;19(1):122. doi: 10.1186/s13000-024-01544-8. [DOI] [PMC free article] [PubMed] [Google Scholar]

[bib19] 19.Ott de Bruin LM, Pico-Knijnenburg I, van Ostaijen-Ten Dam MM, et al. Persistent hypogammaglobulinemia after receiving rituximab post-HSCT is not caused by an intrinsic B cell defect. Int J Mol Sci. 2023;24(21):16012. doi: 10.3390/ijms242116012. [DOI] [PMC free article] [PubMed] [Google Scholar]

[bib20] 20.Marzollo A, Serena T, Mainardi C, et al. Hypogammaglobulinemia in children after hematopoietic stem cell transplantation and rituximab treatment: relevance of B cell subsets. J Pediatr Hematol Oncol. 2023;45(1):e145–e149. doi: 10.1097/MPH.0000000000002582. [DOI] [PubMed] [Google Scholar]

[bib21] 21.Makatsori M, Kiani-Alikhan S, Manson AL, et al. Hypogammaglobulinaemia after rituximab treatment-incidence and outcomes. Qjm. 2014;107(10):821–828. doi: 10.1093/qjmed/hcu094. [DOI] [PubMed] [Google Scholar]

[bib22] 22.Levy R, Mahévas M, Galicier L, et al. Profound symptomatic hypogammaglobulinemia: a rare late complication after rituximab treatment for immune thrombocytopenia. Report of 3 cases and systematic review of the literature. Autoimmun Rev. 2014;13(10):1055–1063. doi: 10.1016/j.autrev.2014.08.036. [DOI] [PubMed] [Google Scholar]

[bib23] 23.Luterbacher F, Bernard F, Baleydier F, Ranza E, Jandus P, Blanchard-Rohner G. Case report: persistent hypogammaglobulinemia more than 10 years after rituximab given post-HSCT. Front Immunol. 2021;12 doi: 10.3389/fimmu.2021.773853. [DOI] [PMC free article] [PubMed] [Google Scholar]

[bib24] 24.Booth C, Gilmour KC, Veys P, et al. X-linked lymphoproliferative disease due to SAP/SH2D1A deficiency: a multicenter study on the manifestations, management and outcome of the disease. Blood. 2011;117(1):53–62. doi: 10.1182/blood-2010-06-284935. [DOI] [PMC free article] [PubMed] [Google Scholar]

[bib25] 25.Pachlopnik Schmid J, Canioni D, Moshous D, et al. Clinical similarities and differences of patients with X-linked lymphoproliferative syndrome type 1 (XLP-1/SAP deficiency) versus type 2 (XLP-2/XIAP deficiency) Blood. 2011;117(5):1522–1529. doi: 10.1182/blood-2010-07-298372. [DOI] [PubMed] [Google Scholar]

[bib26] 26.Vissers LTW, van Ostaijen-Ten Dam MM, Melsen JE, et al. Potential role of B- and NK-cells in the pathogenesis of pediatric aplastic anemia through deep phenotyping. Front Immunol. 2024;15 doi: 10.3389/fimmu.2024.1328175. [DOI] [PMC free article] [PubMed] [Google Scholar]

[bib27] 27.Troilo A, Wehr C, Janowska I, et al. Nonpermissive bone marrow environment impairs early B-cell development in common variable immunodeficiency. Blood. 2020;135(17):1452–1457. doi: 10.1182/blood.2019003855. [DOI] [PMC free article] [PubMed] [Google Scholar]

[bib28] 28.Kraus H, Kaiser S, Aumann K, et al. A feeder-free differentiation system identifies autonomously proliferating B cell precursors in human bone marrow. J Immunol. 2014;192(3):1044–1054. doi: 10.4049/jimmunol.1301815. [DOI] [PubMed] [Google Scholar]

[bib29] 29.Ball LM, Bernardo ME, Roelofs H, et al. Cotransplantation of ex vivo expanded mesenchymal stem cells accelerates lymphocyte recovery and may reduce the risk of graft failure in haploidentical hematopoietic stem-cell transplantation. Blood. 2007;110(7):2764–2767. doi: 10.1182/blood-2007-04-087056. [DOI] [PubMed] [Google Scholar]

[bib30] 30.Lebman DA, Edmiston JS. The role of TGF-beta in growth, differentiation, and maturation of B lymphocytes. Microbes Infect. 1999;1(15):1297–1304. doi: 10.1016/s1286-4579(99)00254-3. [DOI] [PubMed] [Google Scholar]

[bib31] 31.Zehentmeier S, Pereira JP. Cell circuits and niches controlling B cell development. Immunol Rev. 2019;289(1):142–157. doi: 10.1111/imr.12749. [DOI] [PMC free article] [PubMed] [Google Scholar]

[bib32] 32.Dupré L, Andolfi G, Tangye SG, et al. SAP controls the cytolytic activity of CD8+ T cells against EBV-infected cells. Blood. 2005;105(11):4383–4389. doi: 10.1182/blood-2004-08-3269. [DOI] [PubMed] [Google Scholar]

[bib33] 33.Wang N, Satoskar A, Faubion W, et al. The cell surface receptor SLAM controls T cell and macrophage functions. J Exp Med. 2004;199(9):1255–1264. doi: 10.1084/jem.20031835. [DOI] [PMC free article] [PubMed] [Google Scholar]

[bib34] 34.Bottino C, Falco M, Parolini S, et al. NTB-A [correction of GNTB-A], a novel SH2D1A-associated surface molecule contributing to the inability of natural killer cells to kill Epstein-Barr virus-infected B cells in X-linked lymphoproliferative disease. J Exp Med. 2001;194(3):235–246. doi: 10.1084/jem.194.3.235. [DOI] [PMC free article] [PubMed] [Google Scholar]

[bib35] 35.Cannons JL, Tangye SG, Schwartzberg PL. SLAM family receptors and SAP adaptors in immunity. Annu Rev Immunol. 2011;29:665–705. doi: 10.1146/annurev-immunol-030409-101302. [DOI] [PubMed] [Google Scholar]

[bib36] 36.Thiel J, Schmidt FM, Lorenzetti R, et al. Defects in B-lymphopoiesis and B-cell maturation underlie prolonged B-cell depletion in ANCA-associated vasculitis. Ann Rheum Dis. 2024;83(11):1536–1548. doi: 10.1136/ard-2024-225587. [DOI] [PMC free article] [PubMed] [Google Scholar]

[bib37] 37.Ochtrop ML, Goldacker S, May AM, et al. T and B lymphocyte abnormalities in bone marrow biopsies of common variable immunodeficiency. Blood. 2011;118(2):309–318. doi: 10.1182/blood-2010-11-321695. [DOI] [PubMed] [Google Scholar]

[bib39] 38.Marchese A. Endocytic trafficking of chemokine receptors. Curr Opin Cell Biol. 2014;27:72–77. doi: 10.1016/j.ceb.2013.11.011. [DOI] [PMC free article] [PubMed] [Google Scholar]

[bib40] 39.Nakatani T, Sugiyama T, Omatsu Y, Watanabe H, Kondoh G, Nagasawa T. Ebf3+ niche-derived CXCL12 is required for the localization and maintenance of hematopoietic stem cells. Nat Commun. 2023;14(1):6402. doi: 10.1038/s41467-023-42047-2. [DOI] [PMC free article] [PubMed] [Google Scholar]

[bib41] 40.Aoki K, Kurashige M, Ichii M, et al. Identification of CXCL12-abundant reticular cells in human adult bone marrow. Br J Haematol. 2021;193(3):659–668. doi: 10.1111/bjh.17396. [DOI] [PMC free article] [PubMed] [Google Scholar]

[bib42] 41.Gerosa RC, Boettcher S, Kovtonyuk LV, et al. CXCL12-abundant reticular cells are the major source of IL-6 upon LPS stimulation and thereby regulate hematopoiesis. Blood Adv. 2021;5(23):5002–5015. doi: 10.1182/bloodadvances.2021005531. [DOI] [PMC free article] [PubMed] [Google Scholar]

[bib43] 42.Beck TC, Gomes AC, Cyster JG, Pereira JP. CXCR4 and a cell-extrinsic mechanism control immature B lymphocyte egress from bone marrow. J Exp Med. 2014;211(13):2567–2581. doi: 10.1084/jem.20140457. [DOI] [PMC free article] [PubMed] [Google Scholar]

[bib44] 43.Nakashima K, Zhou X, Kunkel G, et al. The novel zinc finger-containing transcription factor osterix is required for osteoblast differentiation and bone formation. Cell. 2002;108(1):17–29. doi: 10.1016/s0092-8674(01)00622-5. [DOI] [PubMed] [Google Scholar]

[bib45] 44.Tormin A, Li O, Brune JC, et al. CD146 expression on primary nonhematopoietic bone marrow stem cells is correlated with in situ localization. Blood. 2011;117(19):5067–5077. doi: 10.1182/blood-2010-08-304287. [DOI] [PMC free article] [PubMed] [Google Scholar]

[bib46] 45.Li H, Ghazanfari R, Zacharaki D, et al. Low/negative expression of PDGFR-α identifies the candidate primary mesenchymal stromal cells in adult human bone marrow. Stem Cell Rep. 2014;3(6):965–974. doi: 10.1016/j.stemcr.2014.09.018. [DOI] [PMC free article] [PubMed] [Google Scholar]

[bib47] 46.Dunleavy K, Hakim F, Kim HK, et al. B-cell recovery following rituximab-based therapy is associated with perturbations in stromal derived factor-1 and granulocyte homeostasis. Blood. 2005;106(3):795–802. doi: 10.1182/blood-2004-08-3198. [DOI] [PMC free article] [PubMed] [Google Scholar]

[bib48] 47.Zehentmeier S, Lim VY, Ma Y, et al. Dysregulated stem cell niches and altered lymphocyte recirculation cause B and T cell lymphopenia in WHIM syndrome. Sci Immunol. 2022;7(75) doi: 10.1126/sciimmunol.abo3170. [DOI] [PMC free article] [PubMed] [Google Scholar]

[bib49] 48.Mc Guire PJ, Cunningham-Rundles C, Ochs H, Diaz GA. Oligoclonality, impaired class switch and B-cell memory responses in WHIM syndrome. Clin Immunol. 2010;135(3):412–421. doi: 10.1016/j.clim.2010.02.006. [DOI] [PMC free article] [PubMed] [Google Scholar]

[bib50] 49.Mueller LP, Luetzkendorf J, Mueller T, Reichelt K, Simon H, Schmoll HJ. Presence of mesenchymal stem cells in human bone marrow after exposure to chemotherapy: evidence of resistance to apoptosis induction. Stem Cells. 2006;24(12):2753–2765. doi: 10.1634/stemcells.2006-0108. [DOI] [PubMed] [Google Scholar]

[bib51] 50.Li J, Law HK, Lau YL, Chan GC. Differential damage and recovery of human mesenchymal stem cells after exposure to chemotherapeutic agents. Br J Haematol. 2004;127(3):326–334. doi: 10.1111/j.1365-2141.2004.05200.x. [DOI] [PubMed] [Google Scholar]

[bib52] 51.Kemp K, Morse R, Sanders K, Hows J, Donaldson C. Alkylating chemotherapeutic agents cyclophosphamide and melphalan cause functional injury to human bone marrow-derived mesenchymal stem cells. Ann Hematol. 2011;90(7):777–789. doi: 10.1007/s00277-010-1141-8. [DOI] [PubMed] [Google Scholar]

[bib55] 52.Meisel R, Heseler K, Nau J, et al. Cytomegalovirus infection impairs immunosuppressive and antimicrobial effector functions of human multipotent mesenchymal stromal cells. Mediators Inflamm. 2014;2014:898630. doi: 10.1155/2014/898630. [DOI] [PMC free article] [PubMed] [Google Scholar]

[bib53] 53.Lévesque JP, Hendy J, Takamatsu Y, Simmons PJ, Bendall LJ. Disruption of the CXCR4/CXCL12 chemotactic interaction during hematopoietic stem cell mobilization induced by GCSF or cyclophosphamide. J Clin Invest. 2003;111(2):187–196. doi: 10.1172/JCI15994. [DOI] [PMC free article] [PubMed] [Google Scholar]

[bib54] 54.Ponomaryov T, Peled A, Petit I, et al. Induction of the chemokine stromal-derived factor-1 following DNA damage improves human stem cell function. J Clin Invest. 2000;106(11):1331–1339. doi: 10.1172/JCI10329. [DOI] [PMC free article] [PubMed] [Google Scholar]

PERMALINK

Acquired nonpermissive BM microenvironment impairs HSC proliferation and maintenance, and B-cell development after HSCT∗†

Melanie de Gier

Jakov Korzhenevich

Franziska M Schmidt

Iga Janowska

Monique M van Ostaijen-ten Dam

Ingrid Pico-Knijnenburg

Davy Cats

Franziska Keller

Petra Hadlova

Mirjam Kunze

Filiz Markfeld-Erol

Kilian Reising

Hailiang Mei

Robbert G M Bredius

Arjan C Lankester

Mirjam van der Burg

Marta Rizzi