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The Journal of Immunology, 2007, 178: 2556-2564.
Copyright © 2007 by The American Association of Immunologists, Inc.

Differential Biological Role of CD3 Chains Revealed by Human Immunodeficiencies1

María J. Recio2,*, Miguel Angel Moreno-Pelayo2,{dagger}, Sara S. Kiliç2,{ddagger}, Alberto C. Guardo*, Ozden Sanal§, Luis M. Allende, Verónica Pérez-Flores*, Angeles Mencía{dagger}, Silvia Modamio-Høybjør{dagger}, Elena Seoane|| and José R. Regueiro3,*

* Inmunología, Facultad de Medicina, Universidad Complutense, Madrid, Spain; {dagger} Unidad de Genética Molecular, Hospital Ramón y Cajal, Madrid, Spain; {ddagger} Pediatric Immunology, Uludag University Medical Faculty, Görükle-Bursa, Turkey; § Immunology Division, Hacettepe University Children’s Hospital, Ankara, Turkey; Inmunología, Hospital 12 de Octubre, Madrid, Spain; and || Inmunobiología Molecular, Hospital Gregorio Marañón, Madrid, Spain


   Abstract
Top
 Abstract
Introduction
 Materials and Methods
 Results
 Discussion
 Disclosures
 References
 
The biological role in vivo of the homologous CD3{gamma} and {delta} invariant chains within the human TCR/CD3 complex is a matter of debate, as murine models do not recapitulate human immunodeficiencies. We have characterized, in a Turkish family, two new patients with complete CD3{gamma} deficiency and SCID symptoms and compared them with three CD3{gamma}-deficient individuals belonging to two families from Turkey and Spain. All tested patients shared similar immunological features such as a partial TCR/CD3 expression defect, mild {alpha}beta and {gamma}{delta} T lymphocytopenia, poor in vitro proliferative responses to Ags and mitogens at diagnosis, and very low TCR rearrangement excision circles and CD45RA+ {alpha}beta T cells. However, intrafamilial and interfamilial clinical variability was observed in patients carrying the same CD3G mutations. Two reached the second or third decade in healthy conditions, whereas the other three showed lethal SCID features with enteropathy early in life. In contrast, all reported human complete CD3{delta} (or CD3{epsilon}) deficiencies are in infants with life-threatening SCID and very severe {alpha}beta and {gamma}{delta} T lymphocytopenia. Thus, the peripheral T lymphocyte pool was comparatively well preserved in human CD3{gamma} deficiencies despite poor thymus output or clinical outcome. We propose a CD3{delta} >> CD3{gamma} hierarchy for the relative impact of their absence on the signaling for T cell production in humans.


   Introduction
Top
 Abstract
 Introduction
Materials and Methods
 Results
 Discussion
 Disclosures
 References
 
Mature {alpha}beta T cells detect peptides on MHC molecules by way of a variable cell surface heterodimer termed the {alpha}beta TCR (1). Before reaching the membrane, variable {alpha}beta TCR heterodimers associate in a preferential sequence with three invariant dimers collectively called CD3 (CD3{delta}{epsilon}, CD3{gamma}{epsilon}, and {zeta}{zeta}) (2). The stoichiometry of the full {alpha}beta TCR/CD3 complex is most likely {alpha}beta/{delta}{epsilon}{gamma}{epsilon}{zeta}{zeta} (3). Once on the cell surface, CD3 proteins translate ligand recognition by {alpha}beta TCR chains into intracellular signals that drive T cell maturation or apoptosis in the thymus, and T cell activation, proliferation, and effector function or anergy/apoptosis in the periphery (4). During early T cell development, some CD3 chains may act alone or assist immature TCR ensembles, such as those containing pre-TCR chains. CD3 chains lack intrinsic enzymatic activity for signal transduction. Rather, they relay on phosphorylation-dependent recruitment and the activation of a number of cytosolic and transmembrane protein tyrosine kinases and adaptors such as Zap-70, Fyn, Lck, TRIM, LAT, SLP76, and SIT (5). The intracellular substrates of each CD3 chain may be different, together with their respective signaling pathways (6).

The complete lack of any of the chains of the invariant dimers causes in humans a group of rare T lymphocyte immunodeficiencies that, in the case of {delta} and {epsilon}, partially resemble their murine models, i.e., severe selective {alpha}beta T lymphocytopenia and absent {alpha}beta TCR/CD3 surface expression, associated with SCID features and early lethality before 3 years of age (for a review, see Ref. 7). In contrast, CD3{gamma} deficiency was reported to allow in humans (8), but not in mice (6, 9), the selection of substantial numbers of polyclonal peripheral T cells that express relatively high levels of functional {alpha}beta TCR/CD3 complexes (10, 11), albeit with an abnormal stoichiometry ({alpha}beta/{delta}{epsilon}{delta}{epsilon}{zeta}{zeta}) and an impaired association to {zeta}{zeta} dimers (11, 12). Two healthy, unrelated individuals lacking CD3{gamma} have been reported to reach their second and third decade, respectively, but the defect was lethal in a sibling with SCID symptoms (13). It is not clear why complete human CD3{gamma} deficiencies are clinically milder and phenotypically leakier for T lymphocyte numbers than in the murine model or complete human CD3{delta} or {epsilon} deficiencies, and further cases can help to settle the matter.

In this study we have characterized the clinical, immunological, and genetic features of two new patients with complete CD3{gamma} deficiency, lethal SCID symptoms, and partial T lymphocytopenia and compare them with three previously reported cases and with other CD3 deficiencies. The results shed light on the relative role of each CD3 chain in TCR/CD3 expression and function in humans.


   Materials and Methods
Top
 Abstract
 Introduction
 Materials and Methods
Results
 Discussion
 Disclosures
 References
 
Case reports

A 7-mo-old boy (subject VI:1 of family 1; Fig. 1A), born in May 2004 to distantly consanguineous parents, suffered chronic diarrhea and pulmonary infections, recurrent otitis media, oral moniliasis, severe diaper dermatitis, and perianal fistula. The immunological workup with BD Biosciences reagents showed 2180 lymphocytes/µl with partial T lymphocytopenia: 43% CD19+, 18% CD16+CD56+CD3–, 21% CD4+CD8–, 18% CD8+CD4–, 14% CD25+, low CD3 (27% with SK7) and TCR{alpha}beta (2% with WT31), but not TCR{gamma}{delta} (7.6% with 11F2, 3.2% also CD8+, 5.6% also CD3+). CD45 and HLA class I expression were both normal as well as IgG, IgM, and IgA (827, 41, and 84 mg/dl, respectively) and IgG1–4 subclasses (606, 69, 34, and <1.4, age-matched ranges 152–951, 18–216, 18–129, and 0–130, respectively). The proliferative responses to PHA and Con A were low compared with the control (6020 vs 43949 and 6504 vs 35760 cpm, respectively). Tested autoantibodies were negative (anti-nuclear Ab, smooth muscle, parietal, thyroglobulin, transglutaminase, and enterocyte). The boy underwent a bone marrow transplant at the age of 13 mo and a retransplant 5 mo later, but he died due to bilateral pneumonia at the age of 20 mo.


Figure 1
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  		FIGURE 1. Pedigrees and haplotype analysis of families 1 (A) and 2 (B) based on the indicated polymorphic markers spanning the CD3GDE region on chromosome 11q23 (inset, shared core haplotype in boldface). Affected status and disease haplotype are depicted in black. The shared mutation is denoted as CD3G:K69X. Homozygous haplotype segregation was observed in patient VI:1 from family 1 and patient IV:4 from family 2. C, Pedigree of a Spanish family (family 3) characterized previously (17 ).

 
His older brother (VI:3) showed very similar symptoms (diarrhea since birth, oral moniliasis, and perianal fistula since 3 mo of age) and immunological findings at 5–6 mo: normal IgG, IgM, and IgA (2010, 229, and 223 mg/dl, respectively); 2473 lymphocytes/µl with partial T lymphocytopenia (26% CD19+, 29% CD16+CD56+CD3–, 18% CD4+CD8–, 11% CD8+CD4–, 2% CD4+CD25+, low CD3 (37% with SK7) and TCR{alpha}beta (1% with WT31), but not TCR{gamma}{delta} (6.7% with 11F2). CD18 expression and neutrophil chemotaxis were normal. Stool cultures were negative. As the diarrhea did not improve, he was referred to Ankara for bone marrow transplantation but died due to sepsis and diarrhea at 9 mo of age.

Molecular genetics

Genomic DNA was extracted following standard methods from peripheral blood samples of patients and unaffected relatives from families 1 and 2 (Fig. 1). Primers were designed to PCR-amplify all exons and the flanking intronic sequences of CD3G (GenBank accession no. NM_000073), CD3D (GenBank accession no. NM_000732), and CD3E (GenBank accession no. NM_000733) genes (see Table I for CD3G). PCR products were purified with the QIAquick PCR purification kit (Qiagen) followed by direct sequencing using the BigDye Terminator V3.1 cycle sequencing ready reaction kit in the ABI PRISM 3100 automatic sequencer (Applied Biosystems).


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  		Table I. Primers used for PCR amplification of human CD3G exons

 
Members of family 1 were screened for the CD3G mutation c.205A->T by digesting exon 3 amplimers with the Tru9I restriction enzyme and running the digestion products in 3% agarose gels.

Subjects from both families were genotyped for microsatellite markers spanning the genetic interval that contains the CD3G, CD3D, and CD3E genes on chromosome 11q23 (D11S898, D11S4111, D11S1356, MICD3E, GDB:179879, D11S1364, D11S925, D11S4089, and D11S1336). The novel MICD3E marker is intragenic to CD3E and was developed by searching for tandem repeats of CA dinucleotide in the sequence contig (GenBank accession no. NT_033899; Homo sapiens genome view, build 35.1) and by designing flanking primers (forward: 5'-ATAGCCCCAAACTTTGCTCAC-3'; reverse: 5'-CATTAACTCTTTGTTACCCCAACTC-3'). The order of markers was established by integrating genetic and physical maps (National Center for Biotechnology Information; http://www.ncbi.nlm.nih.gov) (Table II).


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  		Table II. Haplotype associated with the c.205A->T pathogenic mutation in CD3Ga

 
Abs and flow cytometry

The expression of different surface markers was studied by flow cytometry using standard procedures (10). The following mAb were used: anti-CD3{epsilon} {gamma}/{epsilon}{delta} (UCHT-1) from Immunotech and anti-CD3 (Leu4), anti-CD3 (SK7), anti-CD4 (Leu2a), anti-CD8 (Leu3a), anti-CD45RA (Leu-18), anti-CD45RO (UCHL-1), anti-TCR{alpha}beta (WT31), anti-TCR{gamma}{delta} (11F2), and anti-CD8 (SK1) from BD Biosciences. An anti-CD3{epsilon}{gamma}/{epsilon}{delta} (F101.01) hybridoma supernatant was a gift from Dr. B. Rubin (Centre Hospitalier Universitaire de Purpan, Toulouse, France). The mAb were FITC- or PE-conjugated or purified, and in the latter case a PE-conjugated goat anti-mouse IgG (H+L) from Caltag Laboratories was used as a secondary Ab. Background fluorescence was defined in all cases with an isotype-matched irrelevant mAb from Caltag Laboratories. Briefly, for single- and two-color immunofluorescence, 5 x 105 cells were incubated for 30 min at 4°C with the appropriate mAb in PBS buffer containing 1% FCS. After two washes with PBS, cells were analyzed in an Epics Elite Analyzer cytofluorometer (Coulter Electronics). For comparative stainings we used the mean fluorescence intensity, defined as the average fluorescence value of the corresponding mAb referred to the logarithmic scale of fluorescence intensity along the x-axis of the histograms.

TCR Vbeta repertoire usage (~70% coverage) was determined within CD3+ lymphocytes with the IOTest Beta Mark kit (Beckman Coulter) following the manufacturer’s instructions.

TCR rearrangement excision circles (TRECs)4

Thymic function was evaluated by peripheral blood CD45RA+ T cell numbers or TCR Vbeta usage (see above) or by quantifying TRECs, the highly specific markers for T cells recently produced by the thymus. The {delta} deletion (signal joint) TRECs formed by {delta}Rec-{psi}J{alpha} rearrangement were amplified and quantified in genomic DNA from PBMC by real-time quantitative PCR using a LightCycler system (Roche) as previously described (14). Briefly, fluorescently labeled oligonucleotides were used as reporter probes for standard curve generation and quantification with the manufacturer’s reagent kits, protocols, and software. Serial dilutions of a plasmid clone containing a 375-bp fragment of the {delta} TREC sequence (supplied by D. Douek, National Institute of Allergy and Infectious Diseases, Bethesda, MD) (15) were used to generate standard curves for quantification, and different amounts of a template were used to ensure a linear response. TREC abundance was normalized to cell number by a parallel amplification for a single copy/chromosome gene (beta-globin), a plasmid of which was also used to generate standard curves. Data are expressed as TRECs per 105 cells using mean values from duplicate or triplicate assays for both TRECs and beta-globin, which never varied >10%. Because there are two globin copies per cell, the TRECs content was calculated as [(mean TRECs quantity/mean globin quantity) (2 x 105)]. The detection limit was two copies. Standard stocks were characterized for total cell content and PCR amplicon quantities by repeated measures over time and were saved as a quality assurance measure of all reagents for run-to-run variability.


   Results
Top
 Abstract
 Introduction
 Materials and Methods
 Results
Discussion
 Disclosures
 References
 
Mutation screening and haplotype analysis

Selective partial T lymphocytopenia (T+/–B+NK+ phenotype) in patients VI:1 and VI:3 from family 1 (Fig. 1A), together with low surface CD3 expression, was suggestive of a hereditary recessive form of CD3 deficiency. Therefore, we sequenced all exons and intron/exon boundaries of CD3G, CD3D, and CD3E genes in patient VI:1. No mutations were detected in CD3D and CD3E, but the analysis revealed a homozygous A to T transversion at nucleotide 205 in exon 3 (c.205A->T; Fig. 2, A and B) of CD3G. This mutation results in the substitution of the lysine codon at position 69 by a premature stop codon (p.K69X), which is predicted to truncate the CD3{gamma} protein shortly after the leader peptide and thus cause the pathology. The mutation creates an additional restriction site for the enzyme Tru9I at exon 3. We took advantage of this finding to develop a screening test specific for the mutation (Materials and Methods, and Fig. 2). This test allowed us to follow the segregation of the mutation in the family and showed that it was, as expected, in a heterozygous state in both parents (obligate carriers) and, additionally, in three healthy relatives tested, including the patient’s sister (Fig. 2C).


Figure 2
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  		FIGURE 2. Mutation analysis of family 1. A, Wild-type and mutant sequences (with the Tru9 I restriction site underlined) are localized within CD3G exon 3. The arrows in exon 3 identify PCR primers. LP, EXT, TM, and IC indicate the leader peptide, extracellular, transmembrane, and intracellular domains of the protein, respectively. B, Electropherograms depicting the CD3G exon 3 sequence fragment that contains the mutation obtained from an unaffected subject (wild-type allele), a patient (VI:1, homozygous for the c.205 A->T mutation), and an heterozygous carrier (V:1). C, Tru9 I digestion of exon 3 PCR products (365 bp) from several family members. A wild-type amplimer (such as in IV:7) yielded five fragments of 175, 75, 42, 40, and 33 bp (only the first four are visible in this 3% agarose gel). When the c.205A->T mutation is present, the 175-bp fragment is cleaved in two parts of 138 and 37 bp (not visible). Several family members were heterozygous, whereas only the patient was homozygous for the c.205A->T mutation. MW, molecular mass marker.

 
The mutation had been reported in another family (no. 2; Fig. 1B) described previously (16). Both families 1 and 2 came from the city of Diyarbakir in Southeast Turkey. Genealogic data, as far as available, revealed no relationship between them, but the geographic and ethnic origin of both families suggested the possibility of a common founder. This was investigated by haplotype analysis of microsatellite markers spanning the CD3 region (Fig. 1). These results showed that both families shared a core haplotype associated with the c.205A->T mutation composed of alleles 195, 152, and 121 of markers D11S1356, MICD3E and GDB:179879, respectively, where the last two are in close proximity to the mutation (Table II and Fig. 1). These data suggest the possibility of a common founder allele bearing the c.205A->T mutation causing the CD3{gamma} deficiency in the Turkish population.

Clinical and immunological spectrum of human CD3{gamma} deficiencies

Patient VI:1 (and probably VI:3) from family 1 and patient IV:4 from family 2 therefore carried identical mutations in CD3G. However, their clinical features were disparate: patients VI:1 and VI:3 had clinical signs of SCID starting at 7 or 3 mo of age, respectively, whereas patient IV:4 is presently healthy and well into his teens (Table III). In a previous work, we reported a similar situation in two CD3G compound heterozygous [c.1A->G] plus [c.IVS2-1G->C] brothers belonging to a nonconsanguineous Spanish family (17) (Table III and Fig. 1C). Indeed, one of the siblings died early in life with SCID features (III:3), whereas his older brother (III:2) has reached his third decade in good health. The comparative immunophenotype of the five individuals (Figs. 3 and 4A) did not reflect substantial differences in TCR/CD3 expression levels between them, although the shared selective partial {alpha}beta (and {gamma}{delta}) T lymphocytopenia was more intense early in life (an average 3- to 9-fold in VI:1, VI:3, and III:3 vs 2- to 3-fold in IV:4 and III:2; Fig. 4B). There was no obvious correlation between in vitro functional assays and SCID vs non-SCID condition, because proliferation to several TCR-dependent stimuli was depressed in all patients at diagnosis (Table III). We also measured peripheral blood TRECs, CD45RA+ T cells, and TCR Vbeta usage as a means to estimate thymic output and function (18, 19) (Fig. 5). Unfortunately, early transplantation (patient VI:1) or decease (IV:3, III:3) precluded a more complete analysis of these parameters in SCID patients. Nevertheless, the results indicated that all tested patients had very few peripheral blood thymus emigrants (CD45RA+TREC+) as compared with age-matched controls, although the difference for TRECs was more meaningful in younger patients because TRECs are strongly reduced in normal adults (Fig. 5, A and B). In contrast, the memory (CD45RO+) T cell pool and TCR Vbeta usage were essentially normal by RT-PCR and/or flow cytometry (Ref. 11 ; Fig. 5, A and C). The size and survival of the naive T cell pool is more dependent on thymus output and TCR/MHC interactions, respectively, than the size and survival of the memory T cell pool (20). Thus, the results indicated that the lack of CD3{gamma} impairs thymus production but not the peripheral expansion or accumulation of mature polyclonal T cells, which correlates with the presence of a small thymus in two tested individuals (13). Therefore, the lack of CD3{gamma} can give rise in man to relatively similar immunological features with substantial peripheral T cells despite poor thymus output but with a wide spectrum of clinical symptoms.


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  		Table III. Summary of clinical and immunological data of human CD3{gamma} deficiencies at diagnosis

 

Figure 3
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  		FIGURE 3. Comparative CD3 and CD4 expression between human CD3{gamma}-deficient (dashed line) and control (solid line) T lymphocytes. Similar results were obtained with other CD3 (F101.01), TCR{gamma}{delta} (11F2), and control (CD45) Abs (data not shown). The vertical dotted line in each panel indicates the upper limit of background fluorescence using isotype-matched irrelevant mAb. The numbers in each histogram indicate the mean fluorescence intensity ratio between control and CD3{gamma}-deficient cells, which are useful for comparative purposes between samples analyzed at different times and in different flow cytometers.

 

Figure 4
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  		FIGURE 4. Comparative absolute lymphocyte counts in human complete CD3 deficiencies. Black symbols identify SCID patients. A, Individual total, T, B, and NK lymphocyte counts are plotted as a function of age in comparison with the normal distribution (P5, median, P95) (38 39 ). Due to the CD3 expression defect, {alpha}beta T lymphocytes were defined as CD4+ plus CD8+ or CD8bright, thus excluding most {gamma}{delta} T cells (<8% in all cases). T {gamma}{delta}, B, and NK lymphocytes were defined as 11F2+, CD20+ or surface Ig, and CD56+ or CD16+CD56+, respectively. Thus, {gamma}{delta} T cells may be underestimated due to the TCR/CD3 expression defect. Data from five, eight, and two different individuals or fetuses from three, three, and one unrelated families for CD3{gamma} ({blacktriangleup}{blacktriangledown}{triangleup} {blacksquare}{square}), CD3{delta} (•, • with diagonal bar, and • with horizontal bar) and CD3{epsilon} ({diamondsuit}) deficiencies, respectively (8 40 41 42 43 44 45 46 ). The specific mutations are listed in B. B, Normalized fold reductions in absolute T lymphocyte counts (cell/µl) were calculated from the above data as a ratio relative to the normal median in age-matched controls for each data point for the indicated individuals (top, CD3{gamma} deficiencies, where n is the number of experiments) or disorders (bottom, CD3 deficiencies, where n is the number of different patients sharing the indicated mutations). The control range (labeled C) is P5/median. Subject VI:3 is presumed to carry the p.K69X mutation, hence the brackets. The average normalized reductions in {alpha}beta T lymphocyte counts ranged 2- to 9-fold for CD3{gamma} deficiencies but 100–4600 for CD3{delta} or CD3{epsilon} deficiencies. Reductions in {gamma}{delta} T lymphocyte counts are also shown for comparison, although they are less reliable as explained above.

 

Figure 5
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  		FIGURE 5. Peripheral blood TRECs, CD45RA+ T lymphocytes, and TCR Vbeta usage as a measure of thymus function in human complete CD3{gamma} deficiencies. A, T cell subsets were defined as indicated. Lines indicate normal ranges (P5, median, P95; Ref. 38 ). Symbols identify patients as in Fig. 4. B, The {delta} deletion (signal joint) TRECs were determined as indicated in Materials and Methods. Circles are healthy donors (the gray circle was run with patient samples) and the gray square is the mother of IV: 4 ({gamma}+/– control). For further reference, the thick line represents normal TRECs levels as a function of age (47 ). C, TCR Vbeta mAb binding within CD3+ lymphocytes. Top, Data from patient III:2 (black bars) at 24 years of age are shown in comparison with a normal age-matched control (C; Empty bars) and with the normal range (P10–P90) (48 ). TCR Vbeta usage by RT-PCR at 18 years of age is shown for comparison (reported previously; Ref. 11 ). Note that Vbeta usage may be underestimated due to the TCR/CD3 expression defect. Discordant PCR/mAb binding results may reflect normal age-dependent fluctuations (Vbeta11) or impaired recognition of the mutant TCR/CD3 complex by the mAb involved (Vbeta4) as reported for framework (BMA031; Ref. 11 ) or Vbeta-specific (Vbeta18; Ref. 49 ) mAb recognition of mutant TCR/CD3 complexes (without CD3{gamma} or with TCR{delta} replacing TCR{alpha}, respectively). Bottom, Reactivity patterns for TCR Vbeta2 mAb binding in the indicated patients (thick histograms) at 1, 16, and 24 years of age, respectively, as compared with a control (thin dotted histograms). The profiles are shown as logarithm of relative fluorescence vs cell number. The vertical dotted line in each panel indicates the upper limit of background fluorescence using isotype-matched irrelevant mAbs.

 

   Discussion
Top
 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
Disclosures
 References
 
Founder effect for the c.205A->T CD3G mutation

We have genetically characterized a new familial case of CD3{gamma} deficiency in Turkey. The fact that the CD3G mutation (c.205A->T) is associated with the same CD3-region haplotype in two affected families strongly supports a common founder allele causing CD3{gamma} deficiency in the Turkish population. Therefore the initial molecular diagnosis of Turkish patients with an immunophenotype suggestive of CD3{gamma} deficiency regardless of the accompanying spectrum of clinical symptoms should consider screening for the c.205A->T mutation. The specific screening test developed here may be useful for the genetic diagnosis of this mutation in other laboratories.

Clinical spectrum of CD3{gamma} deficiencies

Despite their relatively similar immunological features, there is clear clinical heterogeneity among human CD3{gamma} deficiencies, ranging from healthy (n = 2, subjects IV:4 of family 2 and III:2 in the Spanish family) to life-threatening SCID cases (n = 2, Spanish subject III:3 and subject VI:1 of family 1; or n = 3, if we consider patient VI:3 of family 1). In contrast, human complete CD3{delta} or CD3{epsilon} deficiencies (n = 8 + 2, respectively) were all SCID infants with essentially no T cells (100- to 4600-fold reduction; Fig. 4B) (21).

Thus, two sets of CD3{gamma}-deficient individuals sharing the same truncating mutations, similar immunological characteristics, and comparable medical care belonged to both clinical extremes. We believe their disparate clinical features reflect the involvement of either environmental or genetic factors (modifying genes) (22, 23). In the case of family 1 (and presumably also family 2, because it shared the same disease core haplotype), CD3D and CD3E were excluded as modifying genes, as they were shown to lack additional pathogenic mutations or nonsynonymous single nucleotide polymorphisms.

Although the functional data are unfortunately incomplete, both SCID and non-SCID patients showed poor PBMC proliferative responses to Ags and mitogens at diagnosis and persistent low TRECs. These results argue against a correlation of poor proliferative responses with the SCID condition. However, we cannot rule out that differences in thymus function (as observed between VI:1 and IV:4; Figs. 4B and 5, B and C) may have caused the disparate clinical outcomes. Collectively, it seems that thymus output is poor in all patients in terms of naive (CD45RA+TREC+) T cells and, thus, functional T cells are scarce early in life. With time, however, as observed in surviving patients the functional polyclonal memory T cells (CD45R0+) accumulate and expand to close to normal numbers. This may explain the normalization of in vitro proliferation assays in non-SCID patients (Table III, footnote g).

Interestingly, a boy with partial CD3{epsilon} deficiency was also healthy and is now in his late teens despite a comparatively more severe TCR/CD3 expression defect (~10-fold compared with 3-fold in CD3{gamma} deficiency). These results support the idea that partial but not complete CD3{epsilon} deficiency has low clinical penetrance (24). Last, the fact that the three patients with SCID features shared severe enteropathy suggests that the CD3{gamma}-chain, and thus T cell function, is particularly important for survival when the intestinal mucosal barrier is breached.

A hierarchy for structural and functional defects in human CD3 deficiencies

The new family confirms that, in sharp contrast to the lack of CD3{delta} or CD3{epsilon}, the complete lack of CD3{gamma} allowed in all cases for the production of substantial numbers of peripheral polyclonal {alpha}beta (and {gamma}{delta}) T cells (Fig. 4 and Ref. 11) expressing relatively high levels of {alpha}beta (and {gamma}{delta}) TCR/CD3 (Fig. 3 and our unpublished results). Thus, the TCR (and the pre-TCR) can be quite functional in humans without CD3{gamma}, but not without CD3{delta} or CD3{epsilon}. This could be due to structural reasons precluding TCR or pre-TCR assembly in the absence of CD3{delta} or CD3{epsilon} as described originally for all {alpha}beta TCR/CD3 chains (25). However, it was later shown in human non-T cells (HeLa) that the expression of incomplete human {alpha}beta TCR/CD3 complexes is possible when either CD3{delta} or CD3{gamma} (but not {epsilon} or any other chain) is absent (26). This suggested a {epsilon} >> {delta} = {gamma} hierarchy for impact of the missing chain on structure. Such a hierarchy could be expected given the fact that {epsilon} is shared by both CD3{gamma}{epsilon} and {delta}{epsilon} dimers in the normal complex (Fig. 6A). Thus, the structural constraint argument holds for CD3{epsilon}–/– but not for CD3{delta}–/– humans. In a lymphoid context (B cell microsomes), {delta}{epsilon} dimers were shown to associate to TCR{alpha} or TCRbeta, whereas {gamma}{epsilon} dimers could only bind to TCRbeta (27), supporting a {delta} > {gamma} structural hierarchy. Also, although both {gamma}–/– and {delta}–/– TCR/CD3 are structurally viable, only the former can signal for T cell selection (and function) in humans despite the high homology of the two chains (66%) (28). A boy with partial CD3{epsilon} deficiency showed substantial T cell numbers and function despite a severe TCR/CD3 expression defect (10-fold) (24), supporting the idea that CD3{epsilon} is structurally critical but less so for signaling when CD3{gamma} and CD3{delta} are present, because even minute amounts of TCR sufficed for T cell selection/expansion in that case. Therefore we believe that there must be some signaling difference between {gamma}–/– and {delta}–/– TCR/CD3 mutant complexes, as proposed in Fig. 6A ({epsilon} >> {delta} ≥ {gamma} hierarchy for impact on structure, but {delta} >> {gamma} for impact on signaling). In fact, mammalian CD3{delta} resembles the chicken CD3{gamma}{delta} precursor more closely than CD3{gamma} (29), suggesting that the latter became specialized later in evolution. Indeed, specialized signaling functions for each CD3 chain have been reported previously (30). The proposed hypothesis could be tested by retroviral transduction of 2A peptide-linked human TCR/CD3 constructs lacking either {gamma} or {delta} and functional analyses of their signaling potential (31).


Figure 6
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  		FIGURE 6. The normal human {alpha}beta TCR/CD3 (or pre-TCR) complex and proposed hierarchy for structural ({epsilon} >> {delta} ≥ {gamma}) and functional ({delta} >> {gamma}) defects in mutant TCR complexes. A, Surface expression is based on published data using transfected HeLa cells (26 ) or human complete {gamma}–/– or partial CD3{epsilon} -deficient T lymphocytes (12 17 24 ). The normal structure follows published models (50 51 ). Faint receptors indicate low expression as detected by flow cytometry. The {zeta}{zeta} dimers were shown to associate poorly to {gamma}–/– TCR/CD3 complexes in human T cells (11 27 ). Promiscuous binding of human {delta}{epsilon} but not {gamma}{epsilon} dimers to TCR{alpha} or TCRbeta is based on in vitro translation data (27 ). Similar structural but not functional constraints apply to mice. The peripheral {alpha}beta T cell pool is indicated by arrows that represent absolute numbers of peripheral blood mature T lymphocytes (Fig. 4; complete {delta}–/– and {epsilon}–/– humans have virtually no T cells). B, Comparative partial (- - -|) or complete (—|) impact of CD3 defects in human and murine T cell development, illustrating that {delta} >> {gamma} in humans, whereas {gamma} >> {delta} in mice for signaling defects at the double-negative (DN)/double-positive (DP) transition ({gamma}–/–, {delta}–/–, and {epsilon}–/– mice have very few or no single-positive (SP) {alpha}beta T cells).

 
Mice are not humans

Gene targeting of CD3 components in mice has shown that the ablation of any CD3 protein essentially blocked T cell development, although at different intrathymic checkpoints and to a different extent (30). Indeed, all CD3 proteins except CD3{delta} are required for T cell selection at the pre-TCR (TCR-beta) checkpoint (double-negative/double-positive transition) with an {epsilon} > {gamma} > {zeta} >> {delta} rank (Fig. 6B). Thus, contrary to what we propose in humans, mice hierarchy for impact on signaling would be {gamma} >> {delta}. However, as observed in humans in vitro, the expression of incomplete murine {alpha}beta TCR/CD3 complexes in non-T cells (3T3) is possible to substantial levels when either CD3{delta} or {gamma} (but not {epsilon} or any other chain) is absent (Ref. 31 and D. Vignali, unpublished observations), supporting the {epsilon} >> {delta} ≥ {gamma} hierarchy for structural constraints. Similar results were obtained in vivo (6, 9, 32). Interestingly, CD3{delta} is also dispensable for mature {gamma}{delta} (but not {alpha}beta) T cell selection and for {gamma}{delta}TCR surface expression in mice, supporting substantial signaling through certain {delta}–/– TCR/CD3 isotypes (33). But the proposed {gamma}{delta}/{gamma}{epsilon}{gamma}{epsilon}{zeta}{zeta} stoichiometry for the murine {gamma}{delta} TCR/CD3 clearly does not hold in humans. {epsilon}–/– humans resemble the mice model, but {gamma}–/– and {delta}–/– human do not (34). For instance, humans but not mice lacking CD3{delta} have no double-positive thymocytes and no {gamma}{delta} T cells, and mice but not humans lacking CD3{gamma} have very few peripheral {alpha}beta or {gamma}{delta} T cells (Figs. 4 and 6B). Also, in a murine {gamma}–/–{delta}–/– double knockout model (35), human but not mouse CD3{delta} restored signaling at the TCR-beta checkpoint (36). Similarly, a human {delta}{epsilon} heterodimer restored pre-TCR functions in {gamma}–/– {delta}–/–, {gamma}–/–, and (partially) {epsilon}–/– mice (37). Thus, human CD3{delta} carries critical signaling cues that can mimic murine CD3{gamma}. Taken together, these results suggest a different role for CD3{gamma} and CD3{delta} in humans and mice in pre-TCR and TCR function during {alpha}beta T cell development. Nevertheless, lymphocyte selection and expansion mechanisms may differ between species, because other immunodeficiencies show also dramatic differences as compared with mice models. For instance, Zap70- or CIITA-deficient humans show normal or substantial numbers of peripheral CD4+ T cells, respectively, and {gamma}c- or Jak3-deficient humans have normal numbers of B cells, whereas their murine counterparts do not. Finally, CD3 gene inactivation in mice, even when kept in pathogen-free facilities, may cause pathological manifestations (enteropathy in CD3{zeta}- or CD3{delta}-deficient mice) that resemble those observed in some humans (CD3{gamma} or {delta} deficiency).


   Acknowledgments
 
We thank Dr. Selcul Sozer (Uludag University Medical Faculty, Görükle-Bursa) for her excellent help with cytometric data; Manuel de la Rosa and Edgar Fernandez-Malavé (Centro de Biologia Molecular Severo Ochoa, Consejo Superior de Investigaciones Cientificas, Madrid) for helpful discussions and critical reading of the manuscript; F. Ihlan Tezcan (Hacettepe University Children’s Hospital, Ankara, Turkey) for clinical data from patient VI:1; Maarten J. M. van Tol (Leiden University Medical Center, The Netherlands), H. Takada (School of Medical Sciences, Kyushu University, Fukuoka, Japan), Dario Vignali (St. Jude Children’s Research Hospital, Memphis, TN), and Edgar Fernandez-Malavé for sharing unpublished data; and Rita Doforno and Sindo Fontán (Hospital la Paz, Madrid), M. A. Muñoz-Fernandez (Hospital Gregorio Marañón), Ramón Rodríguez (Universidad Complutense), and Garbsiñe Roy (Hospital Ramón y Cajal) for laboratory tests.


   Disclosures
Top
 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
 Disclosures
References
 
The authors have no financial conflict of interest.


   Footnotes
 
The costs of publication of this article were defrayed in part by the payment of page charges. This article must therefore be hereby marked advertisement in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.

1 This work was supported by grants from Ministerio de Educación y Ciencia (BFU2005-01738/BMC), Fondo de Investigaciones Sanitarias (05/946), Ministerio de Ciencia y Tecnología (BMC2002-3247), Comunidad Autónoma de Madrid (GR/SAL/0570/2004), and Mutua Madrileña. A.C.G. and V.P.-F. were supported by fellowships from the Universidad Complutense de Madrid and Ministerio de Educación y Ciencia, respectively. Back

2 M.J.R., M.A.M.-P., and S.S.K. are joint first authors. Back

3 Address correspondence and reprint requests to Dr. José R. Regueiro, Departamento de Inmunología, Facultad de Medicina, Universidad Complutense, 28040 Madrid, Spain. E-mail address: regueiro{at}med.ucm.es Back

4 Abbreviation used in this paper: TREC, TCR rearrangement excision circle. Back

Received for publication July 27, 2006. Accepted for publication November 28, 2006.


   References
Top
 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
 Disclosures
 References
 

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