A Novel R140S γc Variant Alters Cellular Distribution, Reduces Surface Expression, and Impairs Cytokine Signaling in Atypical X-SCID

Abstract

The interleukin-2 receptor γ (IL-2Rγ, or γc) is a crucial component of several cytokine receptor complexes. Deficiencies in γc lead to X-linked severe combined immunodeficiency (X-SCID), characterized by recurrent infections due to the absence or dysfunction of T and NK cells, and nonfunctional B cells. Missense variants in the γc extracellular region are linked to atypical X-SCID with normal counts of T, B, and NK cells and less severe symptoms, yet the underlying cellular and molecular mechanisms are not well understood. This study describes a case of atypical X-SCID with a missense variant (c.420 A > T, p.R140S) in the γc extracellular domain, associated with recurrent bacterial, fungal, and viral infections. We found that the R140S variant leads to reduced surface expression and variably affects cytokine receptor signaling. Specifically, STAT5 phosphorylation and proliferation in CD4⁺ T and CD8⁺ T cells are impaired in response to IL-7, a cytokine essential for T cell survival, proliferation and function. Notably, γc^R140S predominantly localizes to the endoplasmic reticulum, in contrast to WT γc, which is found in acidic compartments. Despite this mislocalization, γc^R140S does not trigger unfolded protein responses, and its protein stability and degradation pathways remain unaffected. Nevertheless, cells expressing high levels of γc^R140S exhibited a competitive disadvantage in culture compared to those expressing WT γc, resulting in the enrichment of cells expressing lower levels of γc^R140S. These findings extend our understanding of how mutations in the extracellular domain of γc can lead to reduced protein expression and influence the pathophysiology of atypical X-SCID.

Supplementary Information

The online version contains supplementary material available at 10.1007/s10875-025-01917-8.

Introduction

IL2RG, located on the X chromosome at Xq13, encodes the interleukin 2 receptor subunit γ (IL-2Rγ, CD132), a type I transmembrane glycoprotein expressed on the surface of most lymphocytes. IL-2Rγ plays essential roles in immune function and is referred to as the common γ chain (γc) because it serves as a shared component of the receptor complexes for several critical cytokines, including IL-2, IL-4, IL-7, IL-9, IL-15, and IL-21 [1]. γc is indispensable for assembling the complete cytokine receptor and enhancing its affinity for various cytokines [2]. For instance, in the context of the trimeric IL-2 receptor, the α subunit modulates the affinity and sensitivity of the receptor complex to IL-2, while the β and γc subunits are key to downstream signaling. The α or β subunits individually have low affinity for IL-2, but after binding with the γ subunit, the βγ dimer and αβγ trimer form medium- and high-affinity receptors, respectively [2, 3]. Upon IL-2 binding, IL-2Rβ and IL-2Rγ facilitate the recruitment and activation of JAK1 and JAK3, respectively, leading to phosphorylation of STAT5. The phosphorylated STAT5 dimers then translocate to the nucleus to activate genes critical for the survival, proliferation, differentiation, and apoptosis of T and natural killer (NK) cells, as well as for the regulation of B cell function [4].

Deficiencies in γc result in X-linked severe combined immunodeficiency (X-SCID) in humans, characterized by the absence or dysfunction of T and NK cells, and nonfunctional B cells (T⁻B⁺NK⁻) [5]. As a consequence of these immune cell abnormalities, patients with X-SCID are prone to severe, recurrent infections from a young age, including pneumonia, meningitis, sepsis, and infections caused by viruses, bacteria, fungi, and Pneumocystis jirovecii [6–8]. In recent years, the utilization of prophylactic antibiotics, intravenous immunoglobulin (IVIG), gene therapy via ex vivo transduction of patient CD34⁺ hematopoietic stem cells (HSCs) with WT IL2RG followed by reinfusion, and allogeneic HSC transplantation has significantly improved clinical outcomes for patients with X-SCID [9–12].

Intriguingly, while variants have been identified across various structural domains of γc, not all cases exhibit a severe phenotype. Some individuals with IL2RG variants have normal numbers of T, B, and NK cells (T⁺B⁺NK⁺) and develop late onset and milder infections, indicating atypical X-SCID that may be attributed to a partial loss of γc function [13, 14]. Missense variants in the γc extracellular domain have been linked to such atypical X-SCID [8]. Additionally, T⁺ X-SCID can also be observed in Omenn syndrome or with specific variants such as p.R222S and p.R222C in IL2RG [15, 16]. However, the underlying cellular and molecular mechanisms remain poorly understood. Additionally, despite the variants occurring at the same position, clinical manifestations among patients exhibit considerable variability [6, 14, 17]. Moreover, factors such as chimerism and mosaicism from maternal engraftment or revertant mutations further complicate the diagnosis due to diverse clinical presentations and immune phenotypes [18].

In this paper, we describe a novel point mutation in the extracellular region of γc (c.420 A > T, p.R140S) associated with an atypical X-SCID. This variant leads to decreased γc expression on lymphocytes, impairing JAK-STAT pathway signaling and reducing lymphocyte proliferation. Notably, some T and NK cells exhibited genetic reversions that restored normal γc expression and JAK-STAT pathway signaling. We demonstrate that although this missense variant does not affect γc degradation pathways or half-life, it causes abnormal intracellular distribution. Moreover, cells expressing higher levels of the mutant γc are at a competitive disadvantage and are progressively eliminated during culturing, leading to the predominance of cells with lower levels of γc. Our findings elucidate unique mechanisms by which a missense variant in the extracellular region can result in reduced protein expression and thereby impair its function.

Materials and Methods

Samples

Patient samples and age- and gender-matched healthy controls (HCs) were recruited by the Department of Clinical Immunology at Children’s Hospital of Fudan University. This study was approved by the ethics committee (approval number 2019-048). Both the children and their parents signed informed consent forms prior to the initiation of this study.

Peripheral blood samples were collected in heparin-containing tubes. Peripheral blood mononuclear cells (PBMCs) were isolated via Ficoll gradient centrifugation. Genomic DNA (gDNA) was extracted from peripheral blood, oral mucosa cells, and lymphocyte subpopulations sorted using a CytoFLEX SRT (Beckman, USA) according to the manufacturer’s instructions. Total RNA was extracted from peripheral blood of the patient and HCs using RNAiso Plus (#9109, TaKaRa Bio, Japan), followed by DNase I (#GRT411, TIANGEN Biotech, China) treatment to remove contaminating gDNA. The RNA was then reverse-transcribed using a PrimeScript 1 st strand cDNA Synthesis Kit (#6110A, TaKaRa Bio, Japan).

Genetic Analysis

Protocols for whole-exome sequencing (WES) and analysis were described in our previous study [19]. gDNA from the patient and his family members was enriched to construct a whole-exome library, followed by sequencing on a HiSeq2000 (Illumina, USA). Data analysis was performed by our bioinformatics team. Nucleotide changes observed in aligned reads were identified and reviewed using NextGENe software (SoftGenetics, USA).

The IL2RG variant identified by WES was further validated using Sanger sequencing. Primers (F1-R1, F2-R2) listed in Table S1 were used to amplify the gDNA fragment containing the IL2RG variant via PCR with high-fidelity KOD-plus DNA polymerase (#F1066K, Toyobo, Japan). To confirm the presence and proportion of the reversion, PCR products were purified using a PCR purification kit (#8027011, Dakewe Biotech, China), ligated into the pEASY-Blunt Zero vector (#CB501-01, TransGen Biotech, China), and transformed into competent DH5α cells. The transformed cells were plated and incubated at 37°C overnight, after which individual colonies were picked and sequenced.

Analysis of TCR and BCR Repertoire Diversity

T cell receptor (TCR) and B cell receptor (BCR) repertoires were analyzed by Wuhan SeqHealth Tech Ltd as described [20].

Flow cytometry

Surface Staining

PBMCs from the patient and age- and gender-matched HCs were washed with pre-chilled FACS buffer (PBS containing 2% fetal bovine serum), followed by a 15 min incubation on ice with Human Fc Receptor Blocking Solution (#422302, BioLegend, USA). After washing, PBMCs were incubated on ice for 30 min with various staining mixes (diluted antibodies in FACS buffer) according to the experimental design. The antibodies used are listed in Table S2. After staining, the cells were washed, resuspended in FACS buffer, and analyzed using a FACSVerse flow cytometer (BD Biosciences, USA). Data were analyzed with FlowJo 10.8.1 software (Treestar, USA).

Intracellular Staining

After surface staining, PBMCs were washed with FACS buffer and fixed using IC Fixation Buffer (#00-8222-49, Thermo, USA). The cells were then permeabilized using a pre-diluted Permeabilization Buffer (#00-8333-56, Thermo, USA) and stained with PE α-human CD132 (diluted in permeabilization buffer) for 30 min at room temperature (RT) in the dark.

Phosphoflow

PBMCs (5 × 10⁵) were stimulated with or without 10 ng/mL of IL-21 or IL-7 for 10 min at 37°C, and subsequently fixed with pre-warmed 1×Lyse/Fix Buffer (#558049, BD Biosciences, USA) for 10 min. The cells were centrifuged, washed twice, treated with Fc block, and stained with various surface markers as described in the Surface Staining section. Subsequently, cells were permeabilized with Perm Buffer III (#558050, BD Biosciences, USA) for 30 min at −20°C in the dark, washed three times, and stained with a phospho-STAT antibody or corresponding isotype control for 1 h at RT in the dark. Finally, cells were washed thoroughly and resuspended in FACS buffer for flow cytometric analysis.

Proliferation Assays

PBMCs were washed and resuspended in PBS, shielded from light, and incubated with 3 µM CFSE (#C34554, Invitrogen, USA) for 3 min and 30 s at 37°C. The reaction was stopped by adding 10 mL of PBS containing 5% fetal bovine serum. The labeled cells were washed, resuspended in RPMI1640 medium supplemented with 10% heat-inactivated FBS, antibiotics (100 U/mL penicillin and 100 µg/mL sterptomycin), and 5 × 10⁻⁵ M 2-mercaptoethanol, and seeded at a density of 1 × 10⁵ CFSE-labeled cells per well in a U-bottom 96-well plate, then cultured for 6 days. For B cell proliferation, CFSE-labeled cells were stimulated with combinations of CpG (0.6 µM, #tlrl-2006, Invivogen, France), IL-2 (10 ng/mL, #202-IL-010/CF, R&D Systems, USA), α-CD40 (200 ng/mL, #555587, BD Biosciences, USA), IL-21 (50 ng/mL, #8879-IL-010/CF, R&D Systems, USA), IL-4 (25 ng/mL, #BT-004-010, R&D Systems, USA), and α-IgM (2 µg/mL, #314502, Biolegend, USA). For T cell proliferation, CFSE-labeled cells were stimulated with IL-7 (10 ng/mL, #BT-007-010, R&D Systems, USA) or α-CD3 (2 µg/mL, #317301, Biolegend, USA) + α-CD28 (5 µg/mL, #302901, Biolegend, USA). Following surface staining, CFSE peaks in gated CD4⁺ T, CD8⁺ T, and CD19⁺ B cells were analyzed using flow cytometry.

Establishment of Jurkat Cells Expressing WT or Mutant γc

Primers F3-R3 and F4-R4 were used to amplify the IL2RG coding sequences (CDS) from cDNA of healthy controls and the patient using KOD-plus DNA polymerase through PCR. This yielded the full-length IL2RG CDS excluding the stop codon and incorporating restriction enzyme sites. The PCR products of F3-R3 and the pMX-IRES-EGFP retroviral vector, as well as F4-R4 and the pCDH-EGFP lentiviral vector, were digested with BamHI and NotI, and NheI and AgeI, respectively. These products were then purified and ligated to construct the lentiviral vector pCDH-IL2RG-EGFP, which expresses the γc-EGFP fusion protein, and the retroviral vector pMX-IL2RG-FLAG-IRES-EGFP, which expresses the γc-FLAG fusion protein, respectively. Virus packaging and infection followed previously described methods [21]. After infection, Jurkat cells were cultured for 72 h, and EGFP⁺ cells were sorted using CytoFLEX SRT.

Cell Imaging

Jurkat cells expressing WT or mutant γc were washed with Hank’s Balanced Salt Solution (with Ca²⁺ & Mg²⁺, #C0219, Beyotime, China). Cells were incubated with pre-warmed 60 nM Lyso-Tracker probe (#C34554, Thermo Fisher, USA) that labels acidic compartments including lysosomes and late endosomes in RPMI1640 medium for 1 h in a CO₂ humidified cell incubator. Due to the weak basic nature, the Lyso-Tracker is capable of staining acidic compartments (including lysosomes and late endosomes) within living cells [22, 23]. For endoplasmic reticulum (ER) identification, cells were incubated with pre-warmed, pre-diluted ER-Tracker Red (#C1041S, Beyotime, China) for 30 min. The Golgi apparatus was stained with Golgi-Tracker Red (#C1043, Beyotime, China) for 30 min at 4°C, followed by washing with pre-cooled PRMI1640 medium and incubation with fresh medium at 37°C for another 30 min. Finally, cells were washed again with fresh medium and transferred to glass-bottomed dishes for imaging using a confocal microscope (Leica TCS SP8, Germany).

Immunoblotting

Cells treated with various inhibitors were collected, washed with ice-cool PBS, and then lysed using RIPA lysis buffer containing protease inhibitors. Post centrifugation, the supernatant was mixed with 6×SDS-PAGE Protein Loading Buffer (#P0015F, Beyotime, China), and heated in a metal bath at 95°C for 10 min. Proteins were separated by 4–20% SDS-PAGE and transferred to PVDF membranes. After blocking with 5% non-fat milk in TBS-0.05% Tween at RT, the membranes were incubated with primary antibodies overnight at 4°C, followed by secondary antibodies for 1 h at RT. The membranes were washed with TBST and visualized using a Chemiluminescent Western Blot Imaging System.

Statistical Analysis

Data were analyzed using GraphPad Prism 9 (Version 9.5.1) software. The results are presented as the mean ± standard deviation (SD). The statistical analysis of the data from four groups was performed using one-way ANOVA, and significance was evaluated using the p-value (*p < 0.05, ***p < 0.001).

Results

Clinical Manifestations

The patient, a male, was admitted to Fudan University Children’s Hospital at the age of 10. Beginning at the age of 5, he developed a recurrent cough with yellow purulent sputum and intermittent fever, requiring anti-infective therapy 1–2 times a month. Multiple CT scans of the lungs revealed pulmonary infections and bronchiectasis. The patient underwent multiple bronchoscopy procedures, and the pathogens in the alveolar lavage fluid included Mycoplasma, Haemophilus influenzae, Pseudomonas aeruginosa, and Staphylococcus aureus. An increase in galactomannan (GM) in the alveolar lavage fluid was recorded (4.917), indicating the presence of a fungal infection. Despite treatment with a variety of antibacterial and antifungal drugs, symptoms of cough and expectoration persisted, and the patient eventually exhibited abnormal ventilatory function. In addition to recurring pneumonia, he had a history of thrush, eczema, and viral warts, and an MRI of the head suggested otitis media and sinusitis. The levels of cancer antigen 199 (CA199) and CA125 were significantly elevated, but no neoplastic lesions were found on abdominal CT. At 12 years old, he developed swollen cervical and supraclavicular lymph nodes with tenderness, and a PET-CT suggested reactive hyperplasia. Cytomegalovirus (CMV) was detected in the blood, urine, and sputum, and antiviral therapy with ganciclovir was subsequently initiated. The patient is currently 13 years old, with a height of 152 cm (10th–25th percentile) and a weight of 33.6 kg (below the 10th percentile). He continues to experience recurrent cough and sputum production, requiring intermittent anti-infective therapy. IVIG is not currently being administered. Human leukocyte antigen (HLA) typing has been completed, and hematopoietic stem cell transplantation (HSCT) is planned for the future.

Immunophenotyping

Routine blood examinations and flow cytometric analysis of peripheral blood for immune cell subsets revealed that the patient had normal counts of T, B, and NK cells (Table 1). The patient had an increased proportion of naïve B cells, but significantly reduced proportions of memory B cells and switched memory B cells. He exhibited a decreased proportion of CD4⁺ T cells and an increased proportion of CD8⁺ T cells, resulting in an inverted CD4/CD8 ratio, with a decreased proportion of naïve T cells and a markedly increased proportion of memory T cells (Table 2). Serum levels of immunoglobulins (Ig) G, A, M and E were increased, with IgE levels rising to 5503 KU/L. Allergen testing indicated positive reactions to egg whites (15.9 KU/L) and fungi (2.32 KU/L). Despite receiving hepatitis B vaccination at birth, the patient tested negative for hepatitis B surface antibody at 11 years of age. The TCR and BCR repertoires of the patient did not differ significantly from those of HC (Fig. S1).

Table 1.

Routine blood tests for the patient

Age (year)	10.9	11.0	12.1	12.8	Reference ranges b
White blood cell count (10^9/L)	8.84	10.29	9.77	7.87	4.30–11.30
Neutrophils (10^9/L)	3	4.18	4.36	3.35	1.60–7.80
Lymphocytes (10^9/L)	5.05	4.23	4.46	3.59	1.50–4.60
Lymphocyte subpopulation
CD3+ T cells (/µL)	2131.8	2414.8↑a	2417.6↑	2321.3↑	1325–2276
CD4+ T cells (/µL)	684.86	806.97	706.34	539.91	531–1110
CD8+ T cells (/µL)	1120.87↑	1321.53↑	1415.68↑	1603.02↑	480–1112
CD19+ B cells (/µL)	520.38	768.33↑	393.58	436.81	216–536
CD16+ CD56+ NK cells (/µL)	1183.96↑	567.25	750.49	722.44	246–792
Immunoglobulin
IgG (g/L)	21.6↑	15.6↑	24.1↑	23.3↑	6.09–12.85
IgM (g/L)	4.01↑	1.51	4.02↑	3.2↑	0.67–2.01
IgA (g/L)	5.57↑	3.22↑	3.76↑	4↑	0.52–2.16
IgE (KU/L)	2598.19↑	604.08↑	2137.47↑	1293.57↑	< 100
Serum tumor markers
CA125 (U/mL)	464.2↑	412.2↑	363.2↑	402.7↑	0–35
CA199 (U/mL)	> 1200↑	581.65↑	358.9↑	421.1↑	0–37

^a ↑ indicates a value above the reference range

^b Reference ranges are derived from published literature [24]

Table 2.

Flow cytometric analysis of immune cell subsets in the peripheral blood of the patient

Subset	Marker	Age (year)	Patient	Healthy control (n = 3)	Reference rangesb
CD19+ B (% of lymphocytes)	CD19+	11.0	13.60	8.13/14.50/7.48	9.19–19.48
		12.1	8.42 ↓a	8.82/11.70/10.50
		12.8	7.17 ↓	9.68/13.00/13.50
CD19+ CD20+ B (% of lymphocytes)	CD19+ CD20+	11.0	13.00 ↑a	7.49/6.42/7.26
		12.1	8.19 ↓	8.54/11.40/10.30
		12.8	6.91 ↓	9.52/13.00/12.80
Transitional B (% of CD19+ CD20+ B)	CD19+ CD20+ CD27− CD24hi CD38hi	11.0	4.13	4.64/7.00/0.47	1.75–10.30
		12.1	9.66	2.78/3.04/4.74
		12.8	10.10	1.92/13.60/3.91
Early Transitional B (% of CD19+ CD20+ B)	CD19+ CD20+ CD27− CD21low	11.0	6.94 ↓	9.55/10.90/9.82
		12.1	15.50 ↑	3.34/2.11/4.83
		12.8	11. 30 ↑	2.37/9.97/3.25
Late Transitional B (% of CD19+ CD20+ B)	CD19+ CD20+ CD27− CD21+	11.0	87.60 ↑	70.80/80.40/80.30
		12.1	73.00	66.60/75.70/73.70
		12.8	80.40 ↑	71.13/61.60/72.20
Naïve B (of CD19+ CD20+ B)	CD19+ CD20+ CD27− IgM+ IgDhi	11.0	92.30 ↑	64.30/78.50/73.40	51.84–77.61
		12.1	82.10 ↑	68.10/75.00/72.70
		12.8	90.60 ↑	69.50/68.70/70.90
Memory B (% of CD19+ CD20+ B)	CD19+CD20+CD27+IgD−	11.0	2.01 ↓	16.60/5.21/9.69	8.96–24.09
		12.1	4.71 ↓	17.40/13.80/15.80
		12.8	1.25 ↓	12.20/10.40/12.70
Switched memory B (% of CD19+ CD20+ B)	CD19+ CD20+ CD27+ CD38low IgM− IgD−	11.0	1.56 ↓	13.90/4.45/8.07
		12.1	3.49 ↓	14.00/11.60/14.70
		12.8	1.00 ↓	11.20/9.61/11.50
IgM Memory B (% of CD19+ CD20+ B)	CD19+ CD20+ CD27+ IgMhi	11.0	5.56 ↑	4.71/2.72/4.41
		12.1	3.33 ↓	11.40/6.31/3.95
		12.8	5.24 ↓	7.04/9.29/6.21
IgG Memory B (% of CD19+ CD20+ B)	CD19+ CD20+ CD27+ IgG+	11.0	1.70 ↓	7.37/2.72/5.37
		12.1	1.04 ↓	4.87/1.77/4.00
		12.8	0.54 ↓	4.79/4.77/4.50
Plasmablasts (% of CD19+ B)	CD19+ CD38hi CD27hi CD24−	11.0	1.61	3.66/3.27/0.68	0.70–5.67
		12.1	1.05	2.04/0.98/1.31
		12.8	1.59	1.29/2.19/2.97
DNB (% of CD19+ B)	CD19+ CD27− IgD−	11.0	4.29 ↓	7.57/13.70/9.10
		12.1	6.23 ↑	2.61/3.42/3.42
		12.8	6.83	6.84/4.45/6.24
CD21low B (% of CD19+ B)	CD19+ CD21low CD38low	11.0	4.00 ↓	7.30/6.05/4.69
		12.1	6.73 ↑	3.65/4.01/3.27
		12.8	5.13 ↓	5.94/9.09/8.59
CD3+ T (% of lymphocytes)	CD3+	11.0	57.30	76.10/44.80/48.40	57.10–73.43
		12.1	64.80	60.00/66.30/69.10
		12.8	57.10	65.20/43.50/74.10
CD4+ T (% of lymphocytes)	CD3+ CD4+ CD8−	11.0	16.40 ↓	53.00/23.00/21.80	24.00–38.72
		12.1	19.80 ↓	30.30/45.20/37.30
		12.8	14.00 ↓	44.10/26.80/39.20
CD4 naïve T (% of CD4+ T)	CD3+ CD4+ CCR7+ CD45RA+	11.0	27.60 ↓	73.20/42.40/44.20	39.72–69.59
		12.1	38.40 ↓	47.20/63.50/51.10
		12.8	34.50 ↓	35.80/64.50/72.00
CD4 TCM (% of CD4+ T)	CD3+ CD4+ CCR7+ CD45RA−	11.0	54.70 ↑	20.00/51.90/51.10	24.24–52.73
		12.1	37.10	23.40/22.40/36.80
		12.8	22.60 ↓	30.10/19.90/16.30
CD4 TEM (% of CD4+ T)	CD3+ CD4+ CCR7− CD45RA−	11.0	15.70 ↑	3.52/4.42/3.74	3.40-11.17
		12.1	26.30 ↑	17.60/13.60/12.90
		12.8	37.90 ↑	26.80/13.80/9.62
CD4 TEMRA (% of CD4+ T)	CD3+ CD4+ CCR7− CD45RA+	11.0	1.94 ↑	3.28/1.33/0.99	0.10–1.29
		12.1	1.86 ↑	9.39/0.58/0.51
		12.8	5.06 ↑	7.28/1.87/2.02
Treg (% of CD4+ T)	CD3+ CD4+ CD25hi CD127low	11.0	0.28 ↓	0.52/0.64/0.49
		12.1	0.73 ↓	1.06/0.88/0.98
		12.8	0.13 ↓	0.82/0.38/0.17
CD8 T (% of lymphocytes)	CD3+ CD4− CD8+	11.0	29.40	17.30/16.00/15.20	21.01–33.94
		12.1	34.00 ↑	28.10/19.90/30.10
		12.8	35.10 ↑	14.40/20.20/30.60
CD8 naïve T (% of CD8+ T)	CD3+ CD8+ CCR7+ CD45RA+	11.0	23.60 ↓	81.80/66.30. 63.50	41.41–73.04
		12.1	9.10 ↓	49.20/37.00/67.30
		12.8	11.00 ↓	35.90/50.30/71.70
CD8 TCM (% of CD8+ T)	CD3+ CD8+ CCR7+ CD45RA−	11.0	38.00 ↑	6.65/21.30/23.50	13.21–37.89
		12.1	74.60 ↑	17.90/35.00/19.20
		12.8	68.60 ↑	50.40/29.00/17.10
CD8 TEM (% of CD8+ T)	CD3+ CD8+ CCR7− CD45RA−	11.0	32.30 ↑	5.35/7.45/9.48	1.53–15.39
		12.1	2.12	1.66/2.37/5.26
		12.8	1.75	6.89/1.57/1.87
CD8 TEMRA (% of CD8+ T)	CD3+ CD8+ CCR7− CD45RA+	11.0	5.99	6.24/4.99/3.54	2.01–21.65
		12.1	14.10	31.20/25.70/8.20
		12.8	18.70	6.83/19.20/10.00
Th1 (% of CD3+ T)	CD3+ CD4+ CD45RA− CXCR5− CXCR3+ CCR6− CCR4−	11.0	1.82	1.65/2.85/2.07
		12.1	7.81 ↓	11.60/11.40/12.30
		12.8	1.06 ↑	0.58/0.63/0.23
Th2 (% of CD3+ T)	CD3+ CD4+ CD45RA− CXCR5− CXCR3− CCR6− CCR4+	11.0	8.93 ↑	6.92/5.28/4.18
		12.1	0.60	0.68/0.37/1.00
		12.8	0.80 ↓	2.59/4.40/0.87
Th17 (% of CD3+ T)	CD3+ CD4+ CD45RA− CXCR5− CXCR3− CCR6+ CCR4−	11.0	0.61 ↓	0.95/0.68/0.71
		12.1	2.52	1.76/3.51/3.52
		12.8	1.14 ↓	4.60/2.72/1.41
Tfh (% of CD3+ T)	CD3+ CD4+ CD45RA− CXCR5+ CXCR3− CCR6− CCR4−	11.0	0.64 ↓	1.16/0.94/0.72
		12.1	4.05 ↓	6.05/5.88/5.88
		12.8	3.00	5.34/4.21/2.60
NK (% of lymphocytes)	CD3− CD20− CD14− CD16+ CD56+	11.0	8.28 ↓	5.62/12.10/15.10	10.01–26.98
		12.1	5.18 ↓	7.03/12.00/24.10
		12.8	11.60	8.29/10.00/10.90
CD56bri CD16neg NK (% of lymphocytes)	CD3− CD20− CD14− CD16neg CD56bri	11.0	0.15 ↓	0.46/0.83/0.29
		12.1	0.12 ↓	0.47/0.71/0.69
		12.8	0.042	0.059/0.17/0.013
CD56dim CD16bri NK (% of lymphocytes)	CD3− CD20− CD14− CD16bri CD56dim	11.0	8.58	5.55/11.90/15.20
		12.1	2.57 ↓	6.61/11.10/24.10
		12.8	11.20	6.75/9.72/10.50
CD56dim CD16neg NK (% of lymphocytes)	CD3− CD20− CD14− CD16neg CD56dim	11.0	0.43 ↓	6.36/0.96/1.93
		12.1	0.094 ↓	0.42/0.42/0.52
		12.8	0.69	1.20/1.98/0.48
CD56neg CD16bri NK (% of lymphocytes)	CD3− CD20− CD14− CD16bri CD56neg	11.0	15.20 ↑	0.58/1.14/0.96
		12.1	14.70 ↑	2.22/1.19/1.69
		12.8	4.80 ↑	0.56/1.33/0.44
DCs (% of leukocytes)	CD3− CD20− CD14− HLADR+	11.0	0.84	0.69/1.27/0.50
		12.1	0.87 ↓	2.44/2.06/1.62
		12.8	1.08 ↑	0.81/0.74/1.01
mDCs (% of leukocytes)	CD3− CD20− CD14− HLADR+ CD11c+ CD123−	11.0	0.42	0.33/0.69/0.27
		12.1	0.45	1.61/0.22/1.12
		12.8	0.60	0.53/0.49/0.65
pDCs (% of leukocytes)	CD3− CD20− CD14− HLADR+ CD11c− CD123+	11.0	0.036 ↓	0.13/0.14/0.13
		12.1	0.11	0.046/0.16/0.21
		12.8	0.056	0.040/0.18/0.13
Monocytes (% of leukocytes)	CD3− CD20− CD14+	11.0	1.94	1.85/2.28/1.51
		12.1	3.03	4.20/3.89/1.17
		12.8	5.07 ↑	3.17/4.75/2.23
Classical Monocytes (% of leukocytes)	CD3− CD20− CD14+ CD16− HLADR+	11.0	1.55	1.47/2.16/1.25
		12.1	2.87	4.08/3.84/1.15
		12.8	4.13 ↑	1.30/1.26/1.65
Non-classical Monocytes (% of leukocytes)	CD3− CD20− CD14+ CD16+ HLADR+	11.0	0.33 ↑	0.28/0.082/0.21
		12.1	0.11 ↑	0.084/0.050/0.017
		12.8	0.31 ↑	0.13/0.12/0.11
MDSCs (% of leukocytes)	CD3− CD20− CD14+ CD16− HLADR−	11.0	0.02	0.081/0.025/0.0082
		12.1	0.023 ↑	0.014/0/0
		12.8	0.044 ↓	0.23/1.95/0.10
Granulocytes (% of leukocytes)	CD3− CD20− CD14+ CD56− HLADR−	11.0	41.80 ↑	39.10/31.90/33.60
		12.1	26.40	23.70/24.50/32.60
		12.8	27.80	38.70/31.40/23.40
Basophil (% of leukocytes)	CD3− CD20− CD14− CD56− HLADR− CD123+	11.0	0.57 ↓	2.03/1.27/1.54
		12.1	3.22 ↓	3.94/5.13/5.11
		12.8	1.90	2.15/2.07/0.83

^a ↑ and ↓ indicate values above and below the reference range, respectively, or compared to HCs when the reference range is unavailable

^b Reference ranges are derived from published literature [24]

Genetic Analysis

WES was conducted on genomic DNA extracted from the peripheral blood of the patient and his family members. A missense variant, c.420 A > T (p.R140S), was identified in the patient, whereas his mother and sister were heterozygous for this variant (Fig. 1A). No other known genetic alterations that could account for the patient’s clinical phenotype were identified. Unexpectedly, Sanger sequencing revealed the presence of both IL2RG c.420 A > T and c.420 A > G variants in the patient’s peripheral blood gDNA (Fig. 1B); the latter, a silent mutation, was not detected in the WES data. TA cloning demonstrated that 39 out of 47 sequences (82.98%) from the patient’s peripheral blood gDNA were c.420 A > T, while 8 out of 47 sequences (17.02%) were c.420 A > G (Fig. 1B). However, only c.420 A > T was identified in the gDNA from the patient’s oral mucosa cells (Fig. 1C). Given the localization of the IL2RG gene on the X chromosome, it is probable that the patient inherited the 420T allele from his mother, similar to his sister. The 420G allele likely represents a revertant mutation that occurred in immune cells. Although WES confirmed a normal 46XY karyotype in the patient, maternal engraftment, particularly microchmerism, cannot be completely excluded due to the limited sensitivity of karyotyping. The proportions of the 420T allele and the 420G reversion varied among different lymphocyte subpopulations (Fig. 1D). Specifically, 0 of 20 sequences (0%) in B cells were 420G, while 1 of 11 (9.09%) in CD4⁺ T cells and 3 of 23 (13.04%) in CD8⁺ T cells were 420G. In NK cells, 6 of 14 (42.86%) sequences were 420G. The differential proportions of the 420G revertant among various immune cells likely reflect selective pressure during immune cell development and function. The p.R140S variant is localized in the extracellular domain of γc (Fig. S2), and is predicted to be deleterious by in silico algorithms (Fig. 1E). Based on these findings, we proceeded with a more detailed investigation of this variant.

Fig. 1.

Verification of the variant and identification of a revertant mutation. (A) Family pedigree of the patient. (B) Sequence profiles of PCR products from HC genomic DNA (gDNA), the patient’s gDNA, and monoclonal colonies following ligation of the patient’s PCR products into the T vector. (C) Sanger sequencing of gDNA amplification products from the patient’s oral mucosa cells. (D) Analysis of mutation and reversion rates in different lymphocyte subpopulations. (E) In silico predictions of variant’s deleteriousness

The c.420 A > T (p.R140S) Variant Impairs γc Expression

To evaluate the impact of the R140S amino acid change on γc expression, we used flow cytometry to analyze both cell surface and total (cell surface + intracellular) γc expression in various immune cell subsets before and after α-CD3 + α-CD28 stimulation (Fig. 2). In CD19⁺ B cells, both cell surface and total γc expression levels were lower in the patient compared to HCs, both before and after stimulation (Fig. 2A and B, left column). In the patient’s CD4⁺ and CD8⁺ T cells, a bimodal distribution of γc expression was observed (Fig. 2A and B, 2nd and 3rd columns), consistent with the presence of revertant variant in these cells. The higher γc expression peak was comparable to that of HCs, while the lower peak was notably reduced. Additionally, γc expression levels in NK cells were only slightly lower than those in HCs (Fig. 2A and B, right column), which corresponds with the high proportion of the revertant variant in these cells. Overall, γc protein expression was severely diminished in B cells, substantially reduced in T cells, and moderately decreased in NK cells, correlating with the proportions of the revertant variant in these immune cell subsets.

Fig. 2.

γc expression in the patient’s immune cells. Surface and total (surface + intracellular) γc staining in gated CD19⁺ B cells, CD4⁺ T cells, CD8⁺ T cells, and CD56⁺ NK cells, before (A) and after (B) α-CD3 + α-CD28 stimulation for 3 days. MFI values for γc under each condition are shown

Reduced Phosphorylation of STAT3 and STAT5 in the Patient’s Immune Cells

To determine whether the variant affects IL-21/IL-7/IL-2 receptor signaling, we analyzed the phosphorylation of STAT3 and STAT5 following 10-minute stimulations with IL-21, IL-7 and IL-2, respectively. Before stimulation, the patient’s B cells showed p-STAT3 levels comparable to those of three HCs. However, these cells demonstrated reduced upregulation of p-STAT3 after IL-21 stimulation compared to HCs (Fig. 3A, top panels). Similarly, the patient’s CD19⁻CD4⁻CD8⁻ cells, primarily NK cells and monocytes, also showed p-STAT3 levels comparable to HCs before stimulation, but displayed reduced upregulation after IL-21 stimulation (Fig. 3A, bottom panels). In contrast, both CD4⁺ T and CD8⁺ T cells from the patient had p-STAT3 levels similar to HCs before stimulation and exhibited a similar increase in p-STAT3 levels following IL-21 stimulations (Fig. 3A, 2nd and 3rd rows).

Fig. 3.

Phosphorylation levels of STAT3 and STAT5 following IL-21 and IL-7 stimulation. (A) Levels of phosphorylated STAT3 (p-STAT3) and (B) phosphorylated STAT5 (p-STAT5) in gated CD19⁺ B cells, CD4⁺ T cells, CD8⁺ T cells, and CD19⁻CD4⁻CD8⁻ cells, measured before (red) and after (blue) a 10 min stimulation with 10 ng/mL IL-21 (A) and 10 ng/mL IL-7 (B)

For p-STAT5, B cells from both the patient and HCs showed comparable levels prior to stimulation, with only marginal upregulation following IL-7 exposure (Fig. 3B, top panels). Similarly, the CD19⁻CD4⁻CD8⁻ cells from the patient and HCs had comparable p-STAT5 levels before stimulation. After IL-7 stimulation, a fraction of CD19⁻CD4⁻CD8⁻ cells in HCs demonstrated increased pSTAT5 levels, although the proportion of responsive cells varied among individuals, with HC3 showing the lowest proportion. In contrast, the patient’s CD19⁻CD4⁻CD8⁻ cells displayed minimal upregulation of p-STAT5 levels and lacked a prominent responsive fraction (Fig. 3B, bottom panels). Unlike B cells, CD4⁺ T and CD8⁺ T cells from the patient had p-STAT5 levels comparable to HCs before stimulation but demonstrated significantly impaired upregulation of p-STAT5 levels after IL-7 stimulation (Fig. 3B, 2nd and 3rd rows). In addition to impaired responses to IL-21 and IL-7, the patient’s CD4⁺ T cells, CD8⁺ T cells, and CD19⁻CD4⁻CD8⁻ cells also exhibited significantly reduced pSTAT5 upregulation following IL-2 stimulation (Fig. S3A). Collectively, these results indicate that the R140S variant impairs γc-dependent signaling, resulting in diminished IL-21 responses in B cells and NK cells, and reduced IL-7 and IL-2 signaling in CD4⁺ T cells, CD8⁺ T cells, and NK cells.

Correlation of p-STAT3 and p-STAT5 Levels Induced by IL-21 and IL-7 With Cell Surface γc Expression

The γc^R140S variant exhibited reduced protein expression and diminished levels of p-STAT3 and p-STAT5. To further investigate the relationship between γc expression levels and signaling effects, we analyzed mean fluorescence intensity (MFI) of p-STAT3 and p-STAT5 relative to γc expression. As illustrated in Fig. 4A, p-STAT3 levels correlated with cell surface γc expression following IL-21 stimulation. A similar pattern was observed for p-STAT5 following IL-7 stimulation, as depicted in Fig. 4B. This correlation was particularly pronounced in the patient’s CD4⁺ and CD8⁺ cells, where a fraction of cells that restored γc expression through the revertant mutation displayed relatively higher p-STAT5 levels (Fig. 4B). These findings suggest that the R140S variant primarily impairs γc function by reducing its expression on the cell surface.

Fig. 4.

Correlation of STAT phosphorylation with γc surface levels. (A) p-STAT3 and (B) p-STAT5 levels in gated CD19⁺ B cells, CD4⁺ T cells, CD8⁺ T cells, and CD19⁻CD4⁻CD8⁻ cells following a 10-minute stimulation with 10 ng/mL IL-21 (A) and 10 ng/mL IL-7 (B). The X-axis displays γc levels, and the Y axis shows p-STAT levels

γc Variant Impairs T and B Lymphocyte Proliferation

To assess the impact of the γc^R140S variant on lymphocyte proliferation, we labeled fresh PBMCs with CFSE and analyzed CFSE dilution in gated CD19⁺ B cells, CD4⁺ T cells and CD8⁺ T cells following various stimuli using flow cytometry. A portion of the CD19⁺ B cells from HCs underwent cell division in response to CpG, CpG + IL-2, α-CD40, and α-CD40 + IL-4 + IL-21 (Fig. 5A). In contrast, the patient’s CD19⁺ B cells showed minimal division under the same conditions. Notably, HC2 showed a similarly poor response to α-IgM stimulation as the patient. In response to IL-7, both CD4⁺ and CD8⁺ T cells from HCs underwent multiple rounds of cell division (Fig. 5B); however, only a small fraction of the patient’s CD4⁺ and CD8⁺ T cells exhibited limited proliferation. Nonetheless, the patient’s CD4⁺ T and CD8⁺ T cells displayed substantial cell division comparable to HCs in response to α-CD3 + α-CD28 stimulation (Fig. 5).

Fig. 5.

Analysis of B and T cell proliferation. CFSE-labeled PBMCs from HCs and the patient were cultured with various stimuli for 6 days and analyzed for CFSE dilution in gated CD19⁺ (A), CD4⁺ T (B), and CD8⁺ T (C) cells. The proportions of divided and non-divided cells are shown in the left and right of each panel, respectively

We further used frozen PBMCs of the patient and two HCs and assessed T cell proliferation in response to IL-2 and IL-15, using IL-7 responses as controls. Although cytokine responses in frozen PBMCs were generally weaker than those observed in fresh samples, the patient’s CD4⁺ and CD8⁺ T cells both showed a lower proportion of divided cells in response to IL-7 compared to HCs and moreover lacked the subset of cells that had undergone multiple rounds of division, which was present in HCs (Fig. S3B-C). Similarly, the patient’s CD4⁺ and CD8⁺ T cells exhibited reduced proliferation in response to IL-2 and IL-15 (Fig. S3B-C). These results indicate that the γc^R140S variant impairs B cell proliferation in response to multiple stimuli (except α-IgM) and reduces T cell proliferation in response to IL-7, IL-2, and IL-15, while T cell proliferation induced by α-CD3 + α-CD28 stimulation remained comparable to that of HCs.

Structure of the IL-7-IL-7RA-γc Complex is not Affected by R140S Amino Acid Change

To explore the mechanisms behind the reduced protein expression of the γc^R140S variant, we first employed the AlphaFold3 algorithm to assess the impact of the γc^R140S variant on the structural configuration of the IL-7-IL-7RA-γc complex. As illustrated in Fig. S4A, R140 of γc is located away from the interaction interface with IL-7 or IL-7RA and does not affect hydrogen bond formation between the proteins. The irregularly curled region depicted in Fig. S4B represents a zone where AlphaFold3 exhibits suboptimal predictive performance [25]. Despite this, WT γc and γc^R140S demonstrated a high degree of structural similarity. R140 is located in a curly region between β-Sheets, which does not affect the structure of γc itself (Fig. S4C). Therefore, the R140S amino acid substitution is unlikely to affect the interaction between γc and IL-7RA or the binding of IL-7 to the IL-7 receptor. However, the mutation of the basic amino acid R to the neutral amino acid S may reduce the basicity of γc, potentially resulting in abnormal localization of γc^R140S.

Aberrant Localization of γc^R140S

To investigate the causes of decreased γc expression in the patient’s lymphocytes, we established Jurkat cells expressing γc-EGFP fusion proteins. We observed that WT γc localized in punctate clusters, whereas γc^R140S displayed a disperse distribution (Fig. 6). To further substantiate the localization of WT γc and γc^R140S within living cells, Jurkat cells expressing the γc-EGFP fusion protein were incubated with a series of probes that label acidic compartments, including lysosomes and late endosome, the ER, and the Golgi apparatus [23, 26]. WT γc primarily co-localized with acidic compartments (Fig. 6A) and also showed some co-localization with the ER (Fig. 6B), whereas γc^R140S predominantly co-localized with the ER (Fig. 6B). Neither WT γc nor γc^R140S exhibited significant co-localization with the Golgi (Fig. 6C).

Fig. 6.

Aberrant subcellular localization of γc^R140S. Confocal microscope images of Jurkat cells expressing WT γc-EGFP or γc^R140S-EGFP fusion proteins after incubation with Lyso-tracker (A, magenta), ER-tracker (B, red), Golgi-tracker (C, red). Bright-field images also captured

R140S Variant does not Affect the Half-Life or Degradation Pathways of γc

To investigate whether aberrant protein localization affects the stability and degradation of γc^R140S, we used Jurkat cells recently transduced with retroviruses expressing either WT γc-FLAG or γc^R140S-FLAG fusion proteins and treated them with various inhibitors. The half-life of the proteins, assessed using the protein synthesis inhibitor cycloheximide (CHX), revealed no differences between WT γc and γc^R140S (Fig. S5A). To analyze the degradation pathways of γc, we treated cells with different inhibitors. As depicted in Fig. S5B, the degradation of both WT γc and γc^R140S was inhibited by NH₄Cl, a lysosomal inhibitor, but not by 3-MA (3-methyladenine), an autophagy inhibitor. We then cultured Jurkat cells recently transduced with retroviruses expressing these fusion proteins for four months and repeated the above experiments. This time, we also included MG132, a proteasome inhibitor. Again, we found no differences in the half-life (Fig. S5C) or degradation pathways (Fig. S5D) between WT γc and γc^R140S. These findings suggest that both WT γc and γc^R140S are primarily degraded via the lysosomal pathway, consistent with previous studies [27], and not via autophagy or proteasome pathways. Intriguingly, after four months of culture, the protein levels of γc^R140S were significantly lower than those of WT γc (Fig. S5C, 0 h and Fig. S5D, Medium). These observations prompted us to examine whether γc^R140S might induce or enhance an unfolded protein response (UPR) resulting in apoptosis in cells expressing high levels of γc^R140S. We employed thapsigargin (TG) to induce the UPR in Jurkat cells expressing WT γc-FLAG or γc^R140S-FLAG fusion proteins and analyzed the temporal changes in the protein levels of γc-FLAG, XBP1s, and BIP (Fig. S6A). Upon UPR induction, protein levels of both WT γc and γc^R140S decreased similarly (Fig. S6B), accompanied by comparable increases in the expression of UPR signature proteins, such as XBP1s (Fig. S6C) and BIP (Fig. S6D). Therefore, γc^R140S does not induce a stronger UPR compared to WT γc in Jurkat T cells.

Jurkat Cells Expressing High Levels of γc^R140S Undergo Counterselection During Cell Culture

In Jurkat cells recently transduced with retroviruses expressing WT γc-EGFP or γc^R140S-EGFP fusion proteins, the MFI of EGFP was initially similar but decreased at different rates over time during cell culture. After one month, the MFI for WT γc-EGFP decreased to 75%, whereas that for γc^R140S-EGFP dropped to 54% (Fig. 7A). These findings suggest that Jurkat cells expressing high levels of γc^R140S-EGFP are at a competitive disadvantage in cell culture. Moreover, in Jurkat cells expressing WT γc-FLAG or γc^R140S-FLAG fusion proteins, the ratio of γc^R140S-FLAG relative to WT γc-FLAG protein levels, as measured by immunoblot, progressively declined during culture, from 1.28-fold to 0.45-fold (Fig. 7B), suggesting a counterselection against cells with high levels of γc^R140S expression.

Fig. 7.

Counterselection of Jurkat cells expressing high levels of γc^R140S in cell culture. (A) FACS analysis showing the EGFP MFI in Jurkat cells expressing WT γc-EGFP vs. γc^R140S-EGFP from early (Day 0) to later (Day 31) stages in culture. (B) Comparison of γc^R140S to WT γc expression over time in Jurkat cells expressing WT γc-FLAG vs. γc^R140S-FLAG, analyzed by immunoblot with an α-FLAG antibody. The ratios of γc^R140S-FLAG to WT γc-FLAG protein at each time point (Day 0, Day 10, Day 34, Day 119) are presented

Discussion

In the present study, we report a novel IL2RG missense variant (c.420 A > T, p.R140S) in a patient with atypical X-SCID (T⁺B⁺NK⁺) suffering from recurrent infections. This variant led to decreased surface and total protein expression of γc, which in turn variably impaired STAT phosphorylation in response to IL-21/IL-7/IL-2, resulting in diminished proliferation of T and B lymphocytes. Furthermore, we found that γc^R140S exhibited abnormal subcellular localization, and cells expressing high levels of γc^R140S faced a competitive disadvantage compared to those expressing lower levels.

Through Sanger sequencing, we identified mosaicism in the patient’s immune cells due to a revertant mutation. The proportions of the revertant variant varied, being present in the patient’s T and NK cells but almost absent in B cells. Recent research supports a branching model of development for human NK cells, proposing that T and NK cells may share a common progenitor [28, 29]. In light of this evidence, it is plausible that the reversion originated in a progenitor common to both T and NK cells, occurring after their divergence from B cell lineage.

After IL-21 stimulation, significant reductions in STAT3 phosphorylation were observed in the patient’s B cells and CD19⁻CD4⁻CD8⁻ cells, whereas T cells maintained normal p-STAT3 levels. This indicates that the γc^R140S variant affects cytokine receptor signaling pathways differently across various immune cell subpopulations, potentially due to variations in cytokine receptor expression levels, which require different amounts of γc for optimal expression and function. The preservation of normal p-STAT3 levels in the patient’s CD4⁺ T and CD8⁺ T cells following IL-21 stimulation contrasted with lower p-STAT5 levels after IL-7 stimulation (Fig. 3B). In this case, p-STAT5 levels showed a bimodal peak, mirroring the bimodal expression of γc. Whether IL-7R expression is dependent on γc levels remains unclear. Simultaneous flow cytometry analysis of IL-7R and γc expression will be needed to further investigate their relationship.

A previous study indicated that cells expressing mutant γc maintained normal IL-15R signaling yet exhibited diminished IL-7R signaling in terms of STAT5 phosphorylation [30]. Therefore, different amounts of γc are required for optimal signaling through various cytokine receptors within the same immune cell type. Specifically, IL-7R signaling demands a higher expression of γc compared to IL-15R and IL-21R in CD4⁺ T and CD8⁺ T cells. IL-7 plays a critical role in the development, survival, proliferation, homeostasis, and differentiation of both CD4⁺ T and CD8⁺ T cells [31]. CD4⁺ T cells are essential for initiating and coordinating the immune response, and for activating CD8⁺ T cells and phagocytes through direct cell-cell interactions and the release of cytokines [32]. CD8⁺ T cells are particularly important for direct killing of cells infected by viruses and intracellular pathogens. Therefore, the impaired IL-7R signaling is consistent with the diverse clinical symptoms associated with bacterial, fungal and viral infections.

Peripheral blood analysis showed normal counts of T, B, and NK cells (Table 1), and typical TCR and BCR repertoires compared to HC (Fig. S1), indicating normal lymphocyte development in the patient. However, several features consistent with previous IL2RG variant cases were identified: (1) An inverted CD4/CD8 T-cell ratio with an increased count of CD8⁺ T cells [33–35]. (2) A reduction in naïve T cells and an increase in memory T cells [14, 35, 36]. (3) An increase in naïve B cells, a reduction in memory B cells, and a decrease in class-switched memory B cells, indicating impaired B cell activation and differentiation [13, 15, 37]. (4) Elevated IgE levels [13, 14, 38]. Additionally, the patient in this report had an elevated proportion of CD56^negCD16^bri NK cells.

The predominance of memory over naïve subpopulation indicates that most T cells in the peripheral blood of the patient have already been activated (Table 2). During a normal immune response, naïve T cells are activated by antigen-presenting cells and differentiate into effector T cells that function to eliminate pathogens or infected host cells. Subsequently, the majority of these effector T cells undergo apoptosis, a contraction process that reduces the expanded population of activated T cells to prevent excessive immune activity that could damage tissues and organs [39, 40]. However, a subset of these effector T cells survives and differentiates into memory T cells, including both central and effector memory types [41, 42]. This shift from naïve to predominantly memory T cell subpopulations in the peripheral blood indicates extensive T cell activation and differentiation. In the γc^R140S patient, only a fraction of T cells (those with the revertant mutation) can be normally activated, while the majority of T cells exhibit impaired activation and fail to differentiate into fully functional effector T cells. This leads to inefficient pathogen clearance, resulting in persistent infections and chronic inflammation. Consequently, the patient’s naïve T cells are continuously activated and depleted, while effector memory T cells accumulate, compromising the ability of the immune system to respond effectively to new pathogens.

In contrast to γc^R140S T cells, which exhibit partial activation, γc^R140S B cells fail to activate under in vitro conditions using a variety of stimuli, including CpG, CpG + IL-2, α-CD40, and α-CD40 + IL-4 + IL-21. Consistent with these in vitro findings, there is an increase in naïve B cells and a decrease in memory B cells in the peripheral blood of the γc^R140S patient. The profound defects in B cell activation may be partially attributed to the absence of the revertant that restores normal γc expression. Despite the significant defect in B cell activation and proliferation, the patient peripheral blood contained elevated levels of IgG, IgM, IgA and IgE antibodies. One possible explanation is that although only a small fraction of B cells can be activated, these cells were able to efficiently differentiate into antibody-secreting plasma cells. Further research is necessary to elucidate the underlying mechanisms.

We observed that the R140S substitution led to reduced levels of both cell surface and total γc protein. Additionally, we demonstrated that STAT3 and STAT5 phosphorylation levels, induced by IL-21 and IL-7 stimulation respectively, were correlated with γc levels. Thus, γc^R140S adversely affects immune cell activation due to its reduced protein expression. Atypical X-SCID caused by variants in the extracellular domain of γc has been previously documented, with several cases showing reduced γc expression. One study identified that the P58S substitution disrupts γc plasma membrane targeting due to increased interactions with ER/Golgi and nuclear proteins [43]. In our current study, we found that γc^R140S exhibited a dispersed distribution and co-localized with the ER, in contrast to WT γc, which showed a punctate distribution and predominantly co-localized with acidic compartments. This suggests that γc^R140S may affect proper protein processing or sorting [44]. However, the mislocalization of γc^R140S does not affect its degradation, half-life, or trigger a strong UPR. Therefore, the reduced levels of γc^R140S are not attributable to altered degradation pathways or induction of UPR. We noted that Jurkat cells expressing high levels of γc^R140S exhibited a progressive decrease in protein levels during cell culture. These findings suggest that cells with high γc^R140S expression are at a survival and/or proliferation disadvantage, leading to their counterselection, while those expressing lower levels of γc^R140S were gradually enriched. Based on these observations in Jurkat T cells, it is plausible that the low γc expression in lymphocytes of the γc^R140S patient results from a similar selection process. Our results provide new insights into the mechanisms of reduced protein expression caused by variants in the extracellular domain of γc.

Supplementary Information

Below is the link to the electronic supplementary material.

Author Contributions

L.D. conducted the experiments and analyzed the data. B.S. collected and analyzed the clinical data. L.D. prepared a draft of the manuscript. Q.M., X.M., Y.L., M.Y., and Z.W. contributed to data collection and analysis. W.W., X.H., J.H., and J.S. were responsible for diagnosing and treating the patient, as well as collecting clinical samples. X.W., Z.H., R.Z., X.F., Y.L., and C.L. revised the manuscript. S.C. assisted with the confocal microscope. X.W. and J.Y.W supervised the study. J.Y.W. reviewed and finalized the manuscript.

Funding

This work was supported by the Major Research Plan of the National Natural Science Foundation of China (grant #32330033 to J. Y.W.), the National Natural Science Foundation of China (grant #32270932 to J.Y.W.), and the National Natural Science Foundation for Young Scholar (grant #82202013 to Q.M. and #82402110 to X.M.).

Data Availability

No datasets were generated or analysed during the current study.

Declarations

Consent for Publication

Not applicable.

Ethics Approval

The study received approval from Fudan University Children’s Hospital and was conducted in accordance with the principles outlined in the Declaration of Helsinki. Written informed consent was obtained from the patient and his parents for peripheral blood collection.

Consent To Participate

Written informed consent was obtained from the patient and his parents.

Competing interests

The authors declare no competing interests.