Clinical and demographic information, including age, sex, and familial relationships, was collected from the two siblings with CD19 deficiency and their parents.

Peripheral blood samples were obtained from the CD19-deficient siblings, their unaffected siblings and parents, as well as age-matched healthy controls, following informed consent and in accordance with the guidelines approved by the Ethics Committee of the University Medical Center of Beni Messous.

Serum immunoglobulin levels (IgG, IgA, IgM) were quantified using nephelometry (BN ProSpec^® analyzer; Siemens Healthcare Diagnostics, Marburg, Germany), while serum IgE levels were measured by chemiluminescent immunoassay (CLIA) using the Immulite^® 2000 XPi system (Siemens Healthcare Diagnostics, Germany).

Sera (serum samples) were screened for circulating autoantibodies associated with both organ-specific and non-organ-specific autoimmune diseases. Detection was performed qualitatively by indirect immunofluorescence (IIF) and quantitatively by enzyme immunoassays (EIA), including enzyme-linked immunosorbent assay (ELISA) and chemiluminescent immunoassay (CLIA). For connective tissue disease screening, sera were tested for anti-nuclear antibodies (ANAs) using IIF at a 1:80 dilution on HEp-2 cells (INOVA Diagnostics, San Diego, CA, USA) and for IgM rheumatoid factor. Positive ANA patterns were further titrated by serial dilution and analyzed for specific autoantibodies against Ro/SSA (52 and 60 kDa), La/SSB, Sm, Sm/RNP, Scl-70, dsDNA, and Jo-1 using EIA (Quanta-Lyzer 160/240; CA, USA).

For ulcerative colitis and vasculitis, sera were screened for antineutrophil cytoplasmic antibodies (ANCA) by IIF at a 1:20 dilution on human neutrophil slides (INOVA Diagnostics). Coeliac disease screening involved testing IgA anti-tissue transglutaminase (tTG) antibodies on the Alegria analyzer (Orgentec Diagnostika, Mainz, Germany). Autoantibodies associated with autoimmune hepatitis and primary biliary cirrhosis—including F-actin, liver-kidney microsomal antigen (LKM), soluble liver antigen (SLA), antimitochondrial antibodies (AMA), GP210, and SP100—were measured by ELISA (INOVA Diagnostics, San Diego, CA, USA). For autoimmune thyroiditis, thyroid peroxidase antibodies (TPOAb) and thyroglobulin antibodies (TgAb) were assessed using the Immulite 2000 XPi system (Siemens Healthcare Diagnostics, Germany).

To characterize the major lymphocyte subsets (T, B, and NK cells), and to analyze the T and B cell compartments along with CD19, CD21, and CD81 expression on B cells, the following monoclonal antibodies (mAbs) were used: CD3 (SK7), CD4 (SK3), CD8 (SK1), CD19 (SJ25C1), CD20 (L27), CD21 (B-ly4), CD24 (ML5), CD25 (M-A251), CD31 (WM59), CD38 (HB7), CD45 (HLe-1), CD45RA (L48), CD56 (NCAM16.2), CD81 (JS-81), CD127 (HIL-7R-M21), CCR7 (IA6-2), IgD (IA6-2), TCRαβ (IP26), and FOXP3 (C259D/C7) (all from BD Biosciences); CD16 (3G8), CD45RO (UCHL1), and CD27 (M-T271) (Beckman Coulter); and CXCR5 (51505) (Bio-Techne, R&D Systems Inc.). Detailed antibody panels are provided in Supplementary Table S1 .

For surface marker analysis, 100 µL of whole blood was incubated with monoclonal antibodies for 20 minutes at room temperature in the dark. Red blood cells were lysed, followed by washing. For intracellular staining, cells were fixed, permeabilized, washed, and incubated with the relevant mAbs for 30 minutes before final washing and resuspension in phosphate-buffered saline (PBS). Samples were acquired on a BD FACS Lyric™ flow cytometer (Becton Dickinson, San Jose, CA, USA) and analyzed using BD FACSuite™ Software (v2.1) and FlowJo v10.8 (BD Biosciences).

Gating strategies for T cell subsets included: CD4+ naive T cells (CD4+ CD45RA+ CCR7+), central memory (CD4+ CD45RA– CCR7+), effector memory (CD4+ CD45RA– CCR7–), and terminally differentiated effector memory (TEMRA, CD4+ CD45RA+ CCR7–); CD8+ naive (CD8+ CD45RA+ CCR7+), central memory (CD8+ CD45RA– CCR7+), effector memory (CD8+ CD45RA– CCR7–), and TEMRA (CD8+ CD45RA+ CCR7–) T cells. Regulatory T cells (Tregs) were defined as CD4+ CD25+ CD127 low/– FOXP3+, and T follicular helper (Tfh) cells as CD4+ CD45RO+ CXCR5+.

B cell subsets were classified as follows: naive mature B cells (CD20+ CD27– IgD+), non-class-switched memory B cells (CD20+ CD27+ IgD+), class-switched memory B cells (CD20+ CD27+ IgD–), transitional B cells (CD20+ CD24 hi CD38 hi), plasmablasts (CD20+ CD24– CD38 hi), and autoreactive CD21 low CD38 low B cells.

The surface expression of CD19, CD21, and CD81 was assessed in patients, compared to isotype controls, a healthy unrelated control (13-year-old girl), heterozygous carrier parents, and two non-carrier siblings. Treg and Tfh cell frequencies in the two patients and their parents were also compared to five age- and sex-diverse unrelated healthy controls (three females aged 9, 11, and 33 years; two males aged 9 months and 42 years).

Functionally polarized CD4⁺ T cell subsets—including type 1 helper T (Th1), Th2, and Th17 effector and/or memory cells—are defined by their distinct cytokine secretion profiles. To detect intracellular cytokines, peripheral blood mononuclear cells (PBMCs) were first isolated using a density gradient centrifugation method. PBMCs were stimulated with phorbol 12-myristate 13-acetate (PMA; 50 ng/mL, Sigma-Aldrich) and ionomycin (1 μg/mL, Sigma-Aldrich) in the presence of BD GolgiStop™ (BD Biosciences), and incubated for 5 hours at 37°C with 5% CO₂.

Following stimulation, cells were fixed using BD Cytofix™ Fixation Buffer containing formaldehyde (BD Biosciences) for 15 minutes at room temperature, then washed twice with FACS buffer (containing bovine serum albumin and sodium azide). Permeabilization was performed using BD Perm/Wash™ Buffer (BD Biosciences) for 20 minutes, followed by two additional washes.

Cells were then stained with fluorochrome-conjugated monoclonal antibodies against intracellular cytokines and surface markers, including anti-IFN-γ (clone B27), anti-IL-4 (MP4-25D2), anti-IL-17A (N49-653), CD3 (SK7), and CD4 (SK3) (all from BD Biosciences). Data acquisition was performed using a BD FACS Lyric™ Flow Cytometer and analyzed with BD FACSuite™ Software v1.2 and FlowJo v10.8 (BD Biosciences).

Antibody levels specific to tetanus toxoid, diphtheria toxoid, and pneumococcal capsular polysaccharide (PCP) were quantified using commercial ELISA kits, in accordance with the manufacturers’ instructions. The following kits were used: VaccZyme™ Tetanus Toxoid IgG, VaccZyme™ Diphtheria Toxoid IgG, and VaccZyme™ PCP IgG (all from The Binding Site Group Ltd, Birmingham, UK).

Peripheral blood mononuclear cells (PBMCs) were isolated from heparinized blood samples of patient 1 and an unrelated healthy control (13-year-old female) using Ficoll-Hypaque density gradient centrifugation. A cell suspension of 1 × 10₆ PBMCs/mL in phosphate-buffered saline (PBS) containing 0.1% fetal bovine serum (FBS; Invitrogen, Carlsbad, CA, USA) was labeled with 10 µM carboxyfluorescein succinimidyl ester (CFSE; Invitrogen) for 10 minutes at 37°C in the dark. Following incubation, excess dye was removed by washing.

A total of 3 × 10₅ CFSE-labeled PBMCs were seeded into each well of a 12-well flat-bottom culture plate containing RPMI 1640 medium supplemented with 10% FBS, 100 U/mL penicillin, and 100 µg/mL streptomycin. Cells were stimulated either with 1 µg/mL phytohemagglutinin-M (PHA-M; Sigma-Aldrich) or with anti-CD3/anti-CD28–coated Dynabeads (DB; Gibco). Unstimulated CFSE-labeled PBMCs were cultured under identical conditions as controls.

After incubation, cells were harvested and stained with 7-AAD, anti-CD3, and anti-CD4 antibodies for 30 minutes at room temperature. Following washing, data acquisition was performed using a BD FACSLyric™ flow cytometer (Becton Dickinson, San Jose, CA, USA), and data were analyzed with FlowJo software version 10.8 (BD Biosciences).

Genomic DNA (gDNA) was extracted from whole blood using a previously described protocol (22). Whole-exome sequencing (WES) was conducted on gDNA from the first affected sibling (Patient 1). Identified mutations in the CD19 gene were subsequently confirmed by Sanger sequencing in both affected siblings. Segregation analysis was performed in the parents and unaffected siblings through targeted amplification and sequencing of exon 2 of the CD19 gene.

Both siblings were born to healthy consanguineous Algerian parents ( Figure 2A ). Patient 1 is a 10-year-old girl. Her prenatal and perinatal histories were unremarkable. However, beginning at 8 months of age, she experienced recurrent urinary tract infections and episodes of pyelonephritis, necessitating multiple hospitalizations. No congenital anomalies of the kidneys or urinary tract were identified. From the age of 3 years, she developed recurrent episodes of acute purulent otitis media and sinopulmonary infections, which were managed with antibiotics and did not require hospitalization.

At the age of 6, she presented with an episode of extensive herpetic skin lesions that resolved spontaneously. She also experienced several episodes of mucocutaneous fungal infections, primarily affecting the scalp, skin, and genital area, which responded well to topical antifungal treatment. She received all routine childhood immunizations in accordance with the Algerian national vaccination schedule.

On physical examination, the patient exhibited growth and developmental delays, with a length at -2 standard deviations (SD) and weight at -3 SD. The remainder of the physical examination was unremarkable, including assessments of the lymph nodes, skin, head, heart, neck, oral cavity, and the digestive, urogenital, and respiratory systems.

Patient 2 is a 5-month-old boy who was hospitalized at 5 months of age due to severe pneumonia with pleural effusion and respiratory distress. He was born at term but had a low birth weight (2.1 kg). During the first five months of life, he experienced multiple episodes of upper respiratory tract infections.

On admission, physical examination revealed a high-grade fever (39°C), oxygen saturation of 72% on room air, polypnea at 76 breaths per minute, a hacking cough, and tachycardia with a pulse rate of 140 beats per minute. Growth parameters were significantly below age norms, with a weight of -4 SD and length of -3 SD. Auscultation revealed crackles at the bases of both lungs. Additionally, the patient presented with watery diarrhea. There was no clinical evidence of lymphadenopathy or hepatosplenomegaly.

For Patient 1, initial laboratory evaluation revealed a hemoglobin level of 13.6 g/dL and a white blood cell (WBC) count of 14,200/mm³, with a differential of granulocytes 47.1%, lymphocytes 46.2%, and monocytes 6.7%. Initial immunological investigations ( Table 1 ) showed a moderate decrease in serum IgG at 436 mg/L (reference range: 700–1160 mg/L), while IgA was within normal limits at 144 mg/L (ref: 79–169 mg/L), as were IgM at 42 mg/L (ref: 40–90 mg/L) and IgE at 11.9 IU/mL (ref: <52 IU/mL). Protective antibody titers were markedly reduced, with tetanus and diphtheria toxoid-specific IgG levels being low to undetectable (0.024 IU/mL and 0.001 IU/mL, respectively). Following immunization with Pneumovax^®23 (Pneumococcal Vaccine Polyvalent), specific IgG antibodies against pneumococcal capsular polysaccharides remained low at 30.8 mg/L. IgG subclass analysis showed a selective deficiency of IgG1.

Lymphocyte immunophenotyping by flow cytometry demonstrated an absence of CD19+ B cells (0% of total lymphocytes), while CD20+ B cells were present at 11% (absolute count: 562/mm³). T cell populations included CD3+ T cells at 86.8% (4447/mm³), CD4+ T cells at 40.7% (2086/mm³), CD8+ T cells at 37.6% (1925/mm³), with a CD4/CD8 ratio of 1.08. Natural killer (CD56+) cells accounted for 2.2% (115/mm³). Based on these findings, the patient was started on intravenous immunoglobulin (IVIG) therapy at a dose of 0.4 g/kg every 21 days.

Further in-depth immunophenotyping of T and B cell subsets was performed ( Figures 2B–D ). No significant abnormalities were observed in the frequencies of naïve or effector T cell subsets. Regulatory T cells (Tregs) were within normal limits, while T follicular helper (Tfh) cells were slightly elevated compared to age- and sex-matched healthy controls. In the B cell compartment, despite normal total B cell counts, the frequencies of both switched and non-switched memory B cells and plasmablasts were reduced, with a concomitant increase in naïve B cells. These findings suggest an underlying defect in B cell activation and maturation. The near-absent post-immunization antibody responses against tetanus and diphtheria toxoids, as well as persistently low anti-pneumococcal antibody levels, further support this immunological dysfunction.

For Patient 2, laboratory findings revealed a white blood cell (WBC) count of 15,800/mm³ with a differential of 64% neutrophils, 19.9% lymphocytes, 9% monocytes, and 5.4% eosinophils. Hemoglobin was 11.5 g/dL and platelet count were elevated at 788,000/mm³. Serum immunoglobulin levels showed increased IgG and IgA, while IgM was within normal limits. Renal and hepatic function parameters, as well as serum electrolyte levels, were within normal ranges. C-reactive protein (CRP) was markedly elevated at 24 mg/dL (reference: <6 mg/dL). A chest X-ray revealed right-sided pleuropneumonia.

Lymphocyte subset analysis showed normal percentages but decreased absolute counts for all subsets when compared to age-matched reference values: CD3+ T cells at 79.4% (1482/mm³), CD4+ T cells at 46% (859/mm³), CD8+ T cells at 28.7% (537/mm³), CD20+ B cells at 14.2% (266/mm³), and CD16+CD56+ NK cells at 6.4% (119/mm³). Notably, B cells lacked surface expression of CD19. Post-vaccination antibody responses could not be evaluated, as the patient had only received the BCG vaccine.

Further immunophenotyping of T and B cell compartments revealed an expansion of transitional B cells and a reduction in plasmablasts, while other lymphocyte subpopulations were within normal limits ( Figures 2B–D ). Based on these immunologic findings and the positive family history, a diagnosis of CD19 deficiency was established. The patient was initiated on immunoglobulin replacement therapy at a dose of 0.4 g/kg every three weeks.

To identify the underlying molecular defect, whole exome sequencing (WES) was performed for patient 1, the eldest affected sibling. Bioinformatic analysis revealed a homozygous deletion of a cytosine nucleotide in exon 2 of the CD19 gene at cDNA position 232 (GenBank accession: NM_001770.6), designated as c.232delC. At the genomic level, this corresponds to NC_000016.10:g.28932489del and based on the human genome assembly GRCh37/hg19, the chromosomal position is chr16:28943810delC. This variant was queried in the Genome Aggregation Database (gnomAD) v2.1.1, which is compatible with the hg19 reference genome, and was not reported in either the Exomes or Genomes datasets. The c.232delC mutation is predicted to cause a frameshift resulting in a premature stop codon in exon 2 (p.Leu78Trpfs*52), initiating at codon 78 and leading to a stop codon 52 amino acids downstream ( Figure 3A ). This frameshift is expected to result in nonsense-mediated mRNA decay (NMD) and complete loss of functional CD19 protein expression. According to the American College of Medical Genetics and Genomics (ACMG) criteria, the variant is classified as “pathogenic” (24).

The mutation was confirmed by Sanger sequencing, which verified its segregation with the disease phenotype. Family screening revealed that both parents were heterozygous carriers of the mutation. The symptomatic younger brother (Patient 2) was also found to be homozygous for the same mutation, while the two remaining asymptomatic siblings tested negative for the variant ( Figure 3B ).

The expression of CD19 multimeric complex proteins on B lymphocytes was evaluated by flow cytometry in the affected siblings, their unaffected siblings, parents, and healthy controls ( Figures 4A, B ; Supplementary Figure 1 ). CD19 surface expression was completely absent in both affected siblings, while it was slightly reduced in their heterozygous carrier parents compared to non-carrier siblings and healthy controls. CD21 expression was reduced, but not absent, in both affected patients, whereas CD81 expression remained normal. CD225 expression was not assessed due to lack of validated antibodies.

These findings confirm that the homozygous c.232delC (p.Leu78Trpfs*52) mutation results in a complete loss of CD19 surface expression on B cells in affected individuals, with a partial reduction observed in heterozygous carriers. Moreover, the absence of CD19 appears to impair the expression of CD21, while not affecting CD81, suggesting that CD19 is critical for the stability or surface expression of certain components of the CD19 complex.

In vitro T-lymphocyte proliferation in response to mitogens (PHA) and to anti-CD3/anti-CD28 stimulation was normal ( Figure 4C ). Furthermore, intracellular cytokine staining following CD4+ T cell stimulation showed no abnormalities in the frequencies of CD4+IFNγ+ (Th1), CD4+IL-4+ (Th2), and CD4+IL-17A+ (Th17) cells compared to age- and sex-matched healthy controls ( Figure 4D ). These results indicate that T cell function is preserved, making a primary T cell defect an unlikely cause of the patients’ recurrent infections.

Autoimmune manifestations occur in approximately 30% of individuals with Common Variable Immunodeficiency (CVID) and represent a significant contributor to morbidity and mortality in this population (25–27). Similarly, the presence of autoantibodies has been documented in patients with CD19 deficiency and has been associated with the development of autoimmune features (9, 14, 28). To explore whether the c.232delC (p.Leu78Trpfs*52) mutation was associated with the production of autoreactive antibodies, both affected siblings were screened for organ-specific and non-organ-specific autoantibodies.

Patient 1 showed a speckled pattern of antinuclear antibodies (ANA) on indirect immunofluorescence (IIF) at a low titer (1:160) ( Figure 4E ), while antibodies against extractable nuclear antigens (ENA) and double-stranded DNA (dsDNA) were negative. Notably, anti-F-actin antibodies were slightly elevated (23 U/mL; reference: <20 U/mL), a finding suggestive of a predisposition to type I autoimmune hepatitis. In contrast, Patient 2 exhibited markedly elevated levels of anti-F-actin (39 U/mL) and anti-soluble liver antigen (SLA) antibodies (60 U/mL; reference: <20 U/mL), both serologic markers associated with autoimmune hepatitis. Despite these findings, liver function tests remained within normal ranges in both patients.