Combined Immunodeficiency Caused by a Novel De Novo Gain-of-Function RAC2 Mutation

Ras-related C3 botulinum toxin substrate 2 (RAC2) is a GTPase exclusively expressed in hematopoietic cells that acts as a pivotal regulator of several aspects of cell behavior via various cellular processes. RAC2 undergoes a tightly regulated GTP-binding/GTP-hydrolysis cycle, enabling it to function as a molecular switch. Mutations in RAC2 have been identified in 18 patients with different forms of primary immunodeficiency, ranging from phagocyte defects caused by dominant negative mutations to common variable immunodeficiency resulting from autosomal recessive loss-of-function mutations, or severe combined immunodeficiency due to dominant activating gain-of-function mutations. Here, we describe an 11-year-old girl with combined immunodeficiency presenting with recurrent respiratory infections and bronchiectasis. Immunological investigations revealed low T-cell receptor excision circle/K-deleting recombination excision circles numbers, lymphopenia, and low serum immunoglobulin G. Targeted next-generation sequencing identified a novel heterozygous mutation in RAC2, c.86C > G (p.P29R), located in the highly conserved Switch I domain. The mutation resulted in enhanced reactive oxygen species production, elevated F-actin content, and increased RAC2 protein expression in neutrophils, as well as increased cytokine production and a dysregulated phenotype in T lymphocytes. Furthermore, the dominant activating RAC2 mutation led to accelerated apoptosis with augmented intracellular active caspase 3, impaired actin polarization in lymphocytes and neutrophils, and diminished RAC2 polarization in neutrophils. We present a novel RAC2 gain-of-function mutation with implications for immunodeficiency and linked to functional dysregulation, including abnormal apoptosis and cell polarization arising from altered RAC2 expression. Thus, our findings broaden the spectrum of known RAC2 mutations and their underlying mechanisms.


Introduction
RAC2 is a hematopoietic cell-specific member of the Ras homology (Rho) family of guanosine triphosphatases (GTPases) that functions as a crucial regulator of cell signaling and actin dynamics. RAC2 has roles in numerous important functions, including roles in gene transcription, cell survival, cell adhesion, reactive oxygen species (ROS) production, and cytoskeleton reorganization [1]. Progress in exploring RAC2 function and regulation in mice has facilitated understanding of its physiological roles, for example, regulation of hematopoietic stem cell interaction within the bone marrow niche, erythropoiesis, red blood cell actin dynamics, and phagocyte migration and killing, as well as T and B cell maturation [2]; however, RAC1, which is highly homologous with RAC2, is ubiquitously expressed, and the two proteins are co-expressed in mouse cells of hematopoietic origin. In this context, RAC1 and RAC2 are likely to show functional redundancy, which may obscure their specific roles. Nevertheless, in vivo studies have demonstrated that RAC2 coordinates unique cellular processes [3][4][5][6][7][8].
In recent years, different RAC2 mutations have been identified and reviewed in a small number of patients, revealing further information regarding the vital, nonredundant, and complex functions of RAC2 in humans [9][10][11][12]. A dominant negative monoallelic mutation, p.D57N, classified as causing congenital phagocyte defects, was first described in a male infant in 2000. The infant presented with major symptoms including recurrent progressive soft tissue infections and wound healing failure, mainly resulting from dysregulated neutrophil function, including defective migration, polarization, and reduced superoxide generation [13][14][15]. Another patient with the p.D57N mutation, identified by screening of newborns for severe combined immunodeficiency, initially presented with T-cell lymphopenia, further indicating that RAC2 has complex functional roles [16,17]. Subsequently, two siblings were found to be homozygous for the recessive null RAC2 allele, p.W56X, leading to common variable immunodeficiency, along with T-cell compartment abnormalities. These patient phenotypes were qualitatively distinct from the defects observed in RAC2 −/− mice to some extent, indicating specific roles for RAC2 in human hematopoietic cells [18].
The nucleotide-bound state of the RAC2 GTPase is tightly regulated as a molecular switch that undergoes a GTP-binding/GTP-hydrolysis cycle dominated by guanine nucleotide exchange factors (GEFs) and GTPase activating proteins (GAPs) [19]. Since 2019, several groups have reported germline monoallelic activating gain-of-function mutations in RAC2, including p.G12R, p.P34H, p.Q61R, p.E62K, and p.N92T. These hyperactivating mutations resulted in combined immunodeficiency (CID) or severe combined immunodeficiency (SCID) phenotypes, associated with severe lymphopenia, recurrent sinopulmonary infections, and invasive viral infections, or severe bone marrow hypoplasia and absence of circulating leukocytes [20][21][22][23][24][25], providing crucial insights into the roles of RAC2 in human leukocyte biology and helping to elucidate the significance of RAC2 modification.
To date, 18 patients with diverse RAC2 mutations leading to heterogeneous clinical phenotypes and immunological defects have been reported in the literature. The mechanisms underlying the effects of mutations in this gene, which can affect the immune system in various ways, remain unclear and require elucidation. Herein, we describe the clinical and immunological features of a Chinese patient with a previously unreported heterozygous mutation in RAC2, c.86C > G (p.P29R), including combined immunodeficiency, increased apoptosis, and decreased polarization of actin in lymphocytes and neutrophils.

Patient
The patient enrolled in this study was an 11-year-old girl born to a nonconsanguineous family. Clinical data and blood samples were collected when she first presented to the Children's Hospital of Chongqing Medical University in October 2017. This study was approved by the Medical Ethics Committee of the Children's Hospital of Chongqing Medical University (Approval number: 030/2013). Informed consent was obtained from the patient and her guardians.

Flow Cytometric Analysis of T and B Cells
Circulating follicular helper T (cTfh) cells, Th1 cells, Th2 cells, and Th17 cells, and subsets of cTfh cells were quantified in 50-µL whole blood samples in separate experiments, as described previously [26]. B cell subpopulations were defined as documented elsewhere [27]. Samples were acquired on a FACSCanto II flow cytometer and data were analyzed using FlowJo software.

Quantification of T-Cell Receptor Excision Circles
Real-time quantitative PCR (RT-qPCR) to detect T-cell receptor excision circles (TRECs) and K-deleting recombination excision circles (KRECs) was performed as described previously [30]. Results are reported as numbers of TRECs and KRECs per 10 6 cells or per milliliter of blood.

Analysis of Actin Polymerization
For immunofluorescence staining, CD3 T cells were isolated using an EasySep Human CD3 T Cell positive selection Kit (Stemcell), according to the manufacturer's instructions, then plated on poly-L-lysine-coated slides in the presence of soluble αCD3/αCD28-coated beads or 5 µg/mL low endotoxin azide free (LEAF)-purified anti-CD3 and anti-CD28 antibodies (Biolegend) for 30 min [32]. Isolated neutrophils were plated on slides in the presence of 500 nM PMA (Sigma-Aldrich) [33]. After fixation and blocking, cells were stained with phalloidin (Molecular Probes) at a 1:40 dilution and 49-6-diamidino-2-phenylindole dihydrochloride at 1:10,000 in PBS. Cover slips were imaged on a laser scanning confocal microscope (Nikon, Japan). For analysis of phalloidin mean fluorescence intensity [34], neutrophils were equilibrated for 5 min at 37 °C, followed by agonist stimulation, and cell suspensions transferred to ice-cold BD fixation/permeabilization solution at different time points post-stimulation and incubated on ice for 20 min. F-Actin was stained with FITC-phalloidin for 30 min at 4 °C in darkness. FITC-phalloidin intensity was analyzed using a FACSCanto II flow cytometer.

Analysis of Apoptosis
Apoptosis of peripheral T cells from controls or the patient was assessed with or without 5 µg/mL LEAF-purified anti-CD3 and anti-CD28 antibodies (Biolegend) or 10 µg/mL LEAF-purified FASL (Biolegend) for 48 h [35]. Apoptosis of B cells from controls or the patient was assessed following stimulation with 0.6 μg/mL CpG-ODN (InvivoGen) and 5 μg/mL F(ab)2 goat anti-human IgM (Jackson ImmunoResearch) for 48 h [36]. Apoptosis of isolated neutrophils was examined with or without 0.1 µg/mL LPS (Sigma-Aldrich) or 10 µg/mL LEAF-purified FASL (Biolegend) for 16 h [33]. Cells were stained with Annexin V (to detect apoptosis) and 7-aminoactinomycin D (to detect necrosis). Live cells were identified by the absence of staining for either Annexin V or 7-AAD antibodies (both from BD Biosciences). Apoptosis was also evaluated by detection of intracellular expression of active caspase 3 by flow cytometry analysis [35].

Immunoblotting Analysis of RAC2
Isolated neutrophils were lysed and subjected to sodium dodecyl sulfate/polyacrylamide gel electrophoresis on 12% gels and then transferred to PVDF membranes (Millipore). Blots were then rinsed and blocked in PBS-T containing 5% non-fat dried milk (pH 7.5). After overnight incubation (4 °C) with anti-Rac2 (ab154711, Abcam) and anti-β-actin (Sigma-Aldrich), blots were incubated with horseradish peroxidase-conjugated anti-rabbit IgG (Cell Signaling Technology) and detected using Clarity Western ECL substrate (Bio-Rad).

Statistical Analysis
Data were analyzed using an unpaired two-tailed Student t test. All statistical analyses were conducted using GraphPad Prism 8 software (GraphPad Software, Inc., San Diego, CA). P ≤ 0.05 was considered to indicate a significant difference.

Patient Clinical Characteristics
The patient was from a nonconsanguineous kindred with no significant family history. She first presented with fever and cough at 4 months and was diagnosed with pneumonia, and since had recurrent pneumonia at a frequency of three times each year, which was treated locally with repeated oral and intravenous antibiotics. She also experienced recurrent episodes of hemoptysis from 8 years old, with documented bronchiectasis. Due to the recurrent fever and pneumonia, requiring increasingly longer hospitalization and higher grade antibiotic treatment, our hospital was consulted when she was 10 years old; she weighed 20 kg and her height was 122 cm on admission. On clinical examination, clubbing of her fingers was detected, along with moderate moist rales and hepatosplenomegaly. Laboratory investigations showed constitutive severe lymphopenia and fluctuating granulocytopenia, which was particularly prominent during infection, with large vacuoles visible by light microscopy (Fig. 1a-c), and decreased serum immunoglobulin G (IgG) ( Table 1), with no commonly positive autoantibodies, such as anti-nuclear, anti-neutrophil cytoplasmic, or anti-thyroid peroxidase antibodies. Haemophilus influenzae was repeatedly isolated from sputum on etiological examination, while fungal, cytomegalovirus (CMV), and Epstein-Barr virus (EBV) infections were absent. Chest computer tomography scan revealed extensive bronchiectasis (Fig. 1d). From the beginning of 2019, she began to receive regular intravenous immunoglobulin treatment each month; however, pulmonary infections continued to occur more than five times per year and required hospitalization for combined antibiotic treatment.

Immunophenotype
The patient had severe T, B, and NK cell lymphopenia, with high relative percentages of effector/memory T cells, while a markedly reduced CD4:CD8 ratio, as well as recent thymic and bone marrow emigrants ( Table 2). She also showed diminished proliferative responses to stimulation with mitogens or T-cell receptor (TCR), while cytokine production was elevated ( Fig. S1 and Fig. S2). Further workup revealed extended active cTfh, as well as increased percentages of Th1 and Th1/17-like subsets among non-cTfh and cTfh cells (Fig. 2a, b). Additionally, the patient had a significatively increased percentage of double-negative B cells, along with CD21 low B cells (Fig. 2c). Collectively, these results indicated combined immunodeficiency with a dysregulated phenotype.

A Novel Heterozygous RAC2 Mutation Identified by Gene Sequencing
Sequencing analysis using an NGS-based immunodeficiency panel revealed a novel de novo heterozygous variant in RAC2 (c.86C > G), which was confirmed by Sanger sequencing, and causes a proline to arginine substitution at position 29 (p.P29R) of the RAC2 protein (Fig. 3a, b). Pathogenicity analysis of this novel mutation predicted that it is deleterious ( Table 3). The mutated P29 residue resides within a highly conserved Switch I domain (Fig. 3c), which is vitally important for interactions with GEFs and downstream effectors [37]; however, 3D-modeling structural analysis of the mutant protein using Swiss PdbViewer did not show any obvious destruction of hydrogen bonds or influence on interactions with GDP (data not shown). Nevertheless, data on RAC2 and RAC1 mutations in cancer cells support the hypothesis that this novel mutation is disease causing; RAC1 p.P29S substitutions have been identified in sun-exposed melanomas, and head and neck squamous cell carcinomas, as well as analogous RAC2 p.P29L and p.P29Q mutations [38][39][40], supporting the importance of this proline residue for normal Switch I region function. This variant meets the American College of Medical Genetics (ACMG) criteria for pathogenesis [41], as follows: pathogenic strong (PS1) evidence, as it is a de novo mutation; pathogenic moderate 1 (PM1), due to its location in a well-established functional domain; pathogenic moderate 2 (PM2), as the mutation is not reported in population databases, such as the 1000 Genomes Project; and pathogenic supporting 3 (PP3), since multiple lines of computational evidence support a deleterious effect. Taken together, these findings indicate that the variant is plausibly a novel pathogenic heterozygous RAC2 mutation.     [42]. Neutrophils from RAC2 −/− mice and patients with the RAC2 p.D57N mutation exhibit impaired ROS production in response to fMLP, whereas patients with activating gain-of-function mutations, such as RAC2 p.E62K and p.P34H, show enhanced ROS generation [20,21]. Samples from our patient also showed increased ROS production in response to fMLP and WKYMVM, while levels were normal following PMA stimulation (Fig. 4a), since the latter is independent of RAC2 [20].

Enhanced Neutrophil ROS Production and Activation
These findings were reminiscent of the enhanced ROS production and stimulation-induced apoptosis in BTKdeficient human neutrophils, resulting from increased RAC2 activity in the plasma membrane in a primed state, which has been proposed as responsible for neutropenia and tissue damage [33]. Furthermore, neutrophil activation was quantified by multicolor flow cytometry to detect cell-surface CD11b and CD62L expression, and degranulation was assessed by analysis of cell surface expression   of CD35 (secretory vesicle), CD63 (azurophilic granules), and CD66b (gelatinase granules). Samples from our patient showed increased neutrophil activation, but no obvious degranulation abnormalities in response to different stimuli (Fig. S3).

Impaired Actin Polarization in T Cells and Neutrophils
The activated state of the RAC2 GTPase is tightly regulated, and appropriate inactivation of RAC2 is required for F-actin disassembly and phagocytic cup closure [43]. Elevated F-actin content was detected in neutrophils from our patient (Fig. 4b). Decreased cycling between F-actin and G-actin is expected to result in impaired actin polarization, which was observed in patient T cells and neutrophils (Fig. 5a, b), along with impaired RAC2 polarization in neutrophils (Fig. 5c). The apparent macropinosomes detected in neutrophils from our patient were consistent with the abnormal macropinocytosis reported in patients with RAC2 p.E62K mutations [21]. These abnormalities are established as associated with numerous functions, including cellular migration, intracellular trafficking, and phagocytosis, among others. These results demonstrate that tight regulation of RAC2 activity is pivotal for the control of actin dynamics and actin cytoskeleton reorganization required for physiological cell polarization and macropinocytosis.

Increased Apoptosis in T/B Cells and Neutrophils
The RAC2 p.G12R mutation is proven to abrogate the survival and differentiation capacities of hematopoietic stem and progenitor cells because of consequent defects in cellular and mitochondrial networks [24]. Analogously, transgenic mice expressing the constitutively active RAC2 p.G12V or p.Q61L mutations exhibit increased thymocyte and T lymphocyte apoptosis [44]. Furthermore, the RAC2 p.P34H mutation results in increased apoptosis of unstimulated and TCR-stimulated T cells in vitro and EBV-transformed B cells after serum starvation [20]. Mechanistically, these elegant studies have established preliminary and causative roles for RAC2 in lymphopenia. Similarly, significantly increased apoptosis/necrosis of unstimulated, TCR-stimulated, or FASL-induced T cells, and unstimulated or CPG/F(ab) 2 -stimulated B cells, respectively, was observed in vitro in samples from our patient. In addition, we detected increased apoptosis/necrosis of unstimulated and FASL-induced neutrophils, which could not be rescued by LPS stimulation (Fig. 6a). Furthermore, intracellular active caspase 3 expression was markedly increased in leukocytes from the patient (Fig. 6b). Collectively, our data suggest that RAC2 p.P29R leads to increased apoptosis, partly through influencing caspase 3, which may help to dissect the signaling mechanisms underlying her observed severe peripheral lymphopenia and neutropenia.

Strengthened Expression of RAC2 in Neutrophils and Lymphocytes
The RAC2 mutants, p.G12R, p.P34H, p.Q61R, p.E62K, and p.N92T, along with the RAC1 p.P29S and p.N92I alterations, have been shown to exist preferentially in a GTP-bound state, as a result of rapid transition from the GDP-bound state, ultimately resulting in hyperactivity and combined immunodeficiency [20,21,23,24,[38][39][40]. In contrast, the p.D57N mutation acts in a dominant negative fashion to interfere with RAC1 and RAC2 function, and displayed 10% GTP-binding ability, relative to the wild-type protein [13][14][15]. Intriguingly, expression of the homozygous null p.W56X RAC2 mutation was completely absent in cells transfected with the mutant [18]. As no changes in hydrogen bonds or interactions with GDP were predicted to result from the novel mutation reported here, we investigated its influence on RAC2 expression in our patient, and detected increased mRNA and protein levels in neutrophils and lymphocytes (Fig. 7a-d). Hence, we identified altered expression of RAC2 protein arising from this gain-of-function mutation.

Discussion
In this study, we identified a novel heterozygous RAC2 mutation, c.86C > G (p.P29R), in a Chinese patient diagnosed with combined immunodeficiency. The patient suffered from recurrent respiratory infections and bronchiectasis.    deleterious pathogenic mutation. In parallel, we detected increased cytokine production, skewed T lymphocyte phenotype, accelerated apoptosis with augmented intracellular active caspase 3, and impaired polarization of actin in lymphocytes and neutrophils, as well as impaired polarization of RAC2 in neutrophils. Overall, our findings broaden the spectrum of known RAC2 gain-of-function mutations and their underlying pathological mechanisms. The majority of progress in exploring RAC2 function and regulation has been achieved using model mice. A series of murine studies have demonstrated that RAC2 can control apoptosis and cell growth, as well as survival, of hematopoietic cells [45]. Similarly, the RAC2 p.G12R mutation has proven importance for the survival and differentiation capacity of hematopoietic stem and progenitor cells [24]. As mentioned above, RAC2 p.P34H and the p.P29R mutation identified here severely compromise T and B lymphocyte and neutrophil survival, by increasing apoptosis in response to various stimuli. Notably, severe T-cell and B-cell lymphopenia are associated with p.D57N and p.W56X mutations in patients, respectively, which is distinguished from increased peripheral T and B cells in RAC2 − / − mice [4,46,47]. Together, these findings emphasize the importance of tight regulation of RAC2 in humans; however, the mechanisms underlying RAC2 function in human hematopoietic cells have yet to be clearly elucidated.
Various types of RAC2 mutation, including dominant negative, autosomal recessive loss-of-function, and dominant activating gain-of-function, can result in abnormal cytoskeleton reorganization. Efficient cell motility requires both dynamic and regular F-actin polarization, with pseudopodia extending at the front of the cell and retraction at the rear. This physiological status is maintained by restricting active RAC2 to the front of the cell, a function performed by Wiskott-Aldrich syndrome protein (WASP) through its CRIB motif, which removes active RAC2 from the membrane via endocytosis [48]. Furthermore, recent studies have shown that actin depolymerization is crucial for large particle absorption, a process driven by phosphoinositide 3-kinase (PI3K) and the rapid deactivation of RAC and Cdc42, which is essential for the completion of large particle internalization [49]. An elegant study suggested that RAC activation is sufficient to induce membrane ruffling and macropinocytic cup formation, while subsequent RAC deactivation is required for macropinosome closure and further maturation [50]. These findings are consistent with the association of activating RAC2 mutations with increased macropinocytosis and reduced chemotaxis and polarization. We hypothesize that neutrophilia and decreased ROS production, in apparent contrast with neutropenia and increased ROS production alongside macropinocytosis, could be used to facilitate simple preliminary diagnosis of the nature of RAC2 mutations. RAC2 can exhibit distinctive activities in mouse and human T cells. RAC2 acts on the IFNG promoter to increase IFN-γ production and Th1 differentiation [51], consistent with the findings in our patient of increased T-cell cytokine production and skewing of cTfh and non-cTfh subsets toward the Th1-like phenotype. The aberrant differentiation and function of human T cells may contribute to the observed impaired humoral immune responses and inflammatory tissue lesions. Furthermore, the conserved SwitchIdomain in RAC guanosine triphosphatases binds to the PI3K catalytic subunit, and there is crosstalk between the RAC and PI3K signaling pathways [52]; therefore, it is intuitive that some features and pathophysiological mechanisms will be shared between them, as dysregulated cytokine production and accumulation by senescent CD8 T lymphocytes have been proven in patients with activated PI3Kδ syndrome, in addition to defective NK cell maturation and degranulation, which can be modified in vitro by selective inhibition of p110delta in patients with RAC2 p.P34H mutation [20,53,54]. Notably, inverted CD4/CD8 ratio, decreased TRECs/KRECs, and decreased naïve CD4 and CD8 T cells can occur in patients with dominant negative, autosomal recessive loss-of-function, and dominant activating mutations of RAC2, highlighting the need for further clarification of the unique roles of this molecule in the human hematopoietic system.

Conclusion
In conclusion, we report a Chinese patient with a novel heterozygous activating mutation in RAC2, who presented with recurrent pneumonia, bronchiectasis, hypogammaglobulinemia, and severe lymphopenia. Comprehensive immunological assessment indicated that this novel mutation may affect the proliferation, apoptosis, and polarization of leukocytes. The pathogenesis of immunodeficiency in patients with RAC2 mutations remains poorly characterized, and further detailed research is warranted.