A 45-year-old Caucasian female born to first-cousin parents, with two healthy children and without any relevant family history record of prior diseases, was seen at the service of Dermatology (University of Coimbra, Coimbra, Portugal) in March 2014 because of persistent extensive, skin-colored, exuberant, and disfiguring warts in both feet and hands since the age of 10 years (Figure 1). Warts were refractory to treatment with keratolytic agents, cryosurgery, and excision, with minor improvement after treatment with acitretin in association with topical 50% urea cream. Apart from this, the patient did not describe recurrent infection-related episodes or diseases, except for past medical history of measles and mumps during her infancy and varicella infection during her first pregnancy, which all resolved without complications. Of note, such past history of infections is not rare among the patient age-matched Portuguese population since vaccination for these diseases was introduced in the Portuguese national vaccination program years after she was born (1969): in 1974 for measles, in 1987 for mumps, and in 2004 for varicella. In fact, outbreaks of measles and mumps have been reported in Portugal until the late 80s to mid-90s, with peaks of >10,000 cases per year.
In addition, she referred allergic rhino-conjunctivitis treated with cetirizine and fluticasone, and chronic polyarthralgias in the absence of impaired functionality. Serological studies were negative for (severe) infections, including HIV 1/2, HTLV-1, and syphilis. In turn, she showed IgG antibodies for ubiquitous pathogens including CMV >250 arbitrary units (AU/ml) (positive threshold >6AU/ml) and Epstein-Barr virus VCA = 192 U/ml (positive threshold >20 U/ml) and EBNA = 24 U/ml (positive threshold >20 U/ml) in the absence of serum IgM antibodies for these pathogens (0.06 AU/ml; positive threshold >6AU/ml). Slightly increased serum IgG levels (IgG: 1,430 mg/dl), associated with normal IgA (278 mg/dl), IgM (67 mg/dl), C3 (1.4 g/L), C4 (0.33 g/L), and C1 inhibitor (0.318 g/L) serum levels, were detected. In addition, anti-neutrophil and anti-double-strand DNA autoantibodies were negative, while antinuclear autoantibodies were weakly positive. Screening for immunological alterations by flow cytometry (Supplementary Material) using the EuroFlow Primary Immunodeficiency Orientation Tube (PIDOT) showed an absolute defect of CD4-expressing T-cells (<0.01 cells/mul), with normal total T-cell, CD8+ TCRgammadelta- T-cell, and NK-cell (absolute) numbers, associated with consistently increased B-cell counts vs. age-matched normal reference values. Importantly, (TCRalphabeta+ TCRgammadelta-) DN T-cells were significantly expanded (Table 1). Signs/symptoms associated with primary immunodeficiency other than persistent warts in the feet and hands were not observed either at presentation or during the subsequently 5-year follow-up period. Of note, peripheral blood (PB) monocytes and DCs showed no cell surface expression of CD4.
Since rare CD4 polymorphisms that abrogate reactivity of some monoclonal antibodies (MoAbs) with the CD4 molecule have been described, CD4 expression was evaluated using eight different MoAb clones. These eight CD4 MoAb clones were directed against five different epitopes located in two distinct domains of the CD4 protein (Supplementary Material; Figure 2A), as confirmed in competitive staining inhibition experiments (data not shown), in line with previous data in the literature. Detectable levels of either surface membrane (sm) or intracytoplasmic (cy) CD4 expression were observed in none of the cell lineages that usually express this protein in blood of healthy controls, such as T-cells, monocytes, and DCs (Figure 2B). A lack of CD4 expression was confirmed for both classical T-cells and invariant MAIT and iNKT cells (data not shown). In contrast, the expression of other molecules previously associated with CD4 lymphopenia, such as CD3 and HLA-DR, was normal and comparable to that observed in healthy donors (data not shown). Besides no cellular CD4 protein being detected, normal soluble CD4 levels in plasma were observed in the patient using an ELISA assay with a pair of antibodies directed against the extracellular domains of CD4 (amino acids 26-390; Table 2).
CD4 gene sequencing of patient DNA revealed an isolated homozygous mutation (Figure 3; Supplementary Table 1) in the last bp of the 7-8 intron (NC_000012.12: g6818420 G>A), corresponding to the juxtamembrane domain of the CD4 protein. This alteration was considered by the Variant Effect Predictor Tool (VEP) as a splice acceptor variant with a high impact on CD4 protein transcription. No wild type CD4 DNA sequence was detected based on the analysis of the sequence of the amplicon products obtained after PCR amplification of DNA from the patient. Instead, two truncated forms of CD4 RNA/cDNA were detected. Both truncated forms of CD4 RNA presented with a frameshift deletion starting at the juxtamembrane region at the first bp of exon 8 and a premature stop codon associated with a truncated protein with normal extracellular domains in the absence of the anchoring domain to the membrane (Figure 3). The first frameshift deletion (NM_000616: c.1157_1278del) consisted of a complete deletion of exon 8 (122 pb), resulting in a 399-amino-acid protein. In turn, the second frameshift deletion produced a 430-amino-acid protein because of (only) a 29 bp deletion (NM_000616: c.1157_1185del) (Figure 3) that ended just before a 5 bp combination (TGCAG), homologous to the sequence observed at the end of the 7-8 intron (Figure 3).
In order to investigate the origin of the mutation, DNA from four first-degree patient relatives (all asymptomatic) was also analyzed. Thus, the mutation identified in the patient (NC_000012.12: g6818420 G>A) was also found in heterozygosis in DNA from each of her two children and each of her parents. In contrast, her brother's DNA only showed wild type CD4 gene sequences. Further cDNA sequencing from her children and her parents revealed two CD4 mRNA sequences, one carrying the large deletion observed in the patient (NM_000616: c.1156_1278del) and another with the unmutated (wild type) CD4 allele sequence (Figure 3). These results are fully consistent with a germinal mutation in the patient inherited from her parents and transmitted to her children. In contrast, in the patient's brother, both alleles were found to be wild type. Analysis of CD4 protein levels by flow cytometry confirmed that, although normal CD4+ T-cell counts were observed in the patient's children and parents (Table 3), the amount of expression of the CD4 protein on CD4+ T-cells, monocytes, and pDCs was reduced to around half when compared to healthy controls (n = 3) and the patient's brother, stained in parallel (Supplementary Table 2).
As described above, a lack of CD4 expression on patient T-cells was associated with abnormally expanded DN TCRgammadelta- TCRalphabeta+ T-cell counts in PB (>30% of all T-cells; Table 1), compared to age-matched normal reference values. Based on CD27, CD45RA, and CCR7 expression, these DN T-cells showed a similar distribution per maturation stage to that of CD4+ T-cells from age-matched healthy controls, both in relative and absolute numbers: (i) naive: 27% and 240 cells/mu (6-66% and 83-676 cells/mul), (ii) central memory: 66% and 587 cells/mul (18-71% and 235-589 cells/mul), and (iii) effector memory: 7% and 62 cells/mul (2-66% and 14-221 cells/mul). In addition, expanded DN T-cells showed a polyclonal TCRVbeta repertoire with a distribution per TCR family evaluated, fully consistent with reference TCRVbeta repertoire values observed in HLA class II-restricted CD4+ T-cells from healthy donors (Supplementary Figure 1). More extended phenotypic analysis of central/effector memory DN T-cells from the patient showed Treg and Th surrogate marker expression profiles for Tregs, TFH, Th1, Th2, Th17, and Th1/Th17 CD4+ helper T-cell to be present at frequencies similar to those observed for normal CD4+ T-cells (Table 1) from age-matched healthy donors (Botafogo et al., submitted).
Short-term in vitro stimulation, for 4 and 6 h, showed that the expanded DN T-cells were capable of producing cytokines from the main Th patterns (e.g., IFNgamma, IL-4/IL-5, and IL-17A/IL-17F) at frequencies similar to CD4+ helper T-cells from age-matched healthy controls, when either polyclonal (i.e., PMA) or antigen-specific (i.e., CMV) stimuli that required antigen presentation were used (Table 4). In contrast, the percentage of DN T-cells expressing markers of cytotoxic T-cells (cyGranzyme B or cyPerforin) was decreased below <1% (normal age-matched range of <1-21% of DN T-cells).
Normal total CD8+ T-cell, TCRgammadelta+ T-cell, and NK-cell counts were found in the patient blood, although the latter two were increased in some of the monitoring time points tested during the 5-year follow-up period. In contrast, consistently increased naive CD8+ T-cell counts (Table 1) vs. age-matched reference values, associated with normal central and effector memory CD8+ T-cell numbers, were found in the patient's blood at different time points. Expression of cytolytic enzymes (cyGranzyme B and cyPerforin) was detected in CD8+ T-cells (13 and 9% of CD8+ T-cells) and NK-cells (>99%) at normal values (15-65% and 10-53% of CD8+ T-cells, respectively; >99% of NK-cells). In addition, CD8+ TCRalphabeta+ T-cells also showed a normal polyclonal TCRVbeta repertoire (vs. normal age-matched reference values, shown in Supplementary Figure 1) and a normal cytokine production profile in response to PMA and CMV (Table 4).
Detailed dissection of the PB B-cell compartment from the patient showed persistently increased total B-cell counts due to increased naive (CD21+) B-cells, IgG1-3 and IgA1 memory B-cells, and IgA1 plasmablasts, with normal immature/transitional B-cell numbers (Table 1), in line with the observed higher serum IgG levels.
Analysis of circulating PB monocytes based on expression of CD14 (LPS receptor) and CD16 (low-affinity Fc IgG receptor) showed normal absolute counts for all subsets (Table 1), including: (1) CD14+ CD16- classical monocytes (cMo), (2) CD14+ CD16+ intermediate monocytes (iMo), and (3) CD14- CD16+ non-classical monocytes (ncMo). Further dissection of cMo and ncMo based on the expression of the CD62L and SLAN selectins, respectively, did not show significant differences vs. normal age-matched reference values. In addition, CD4-negative monocytes and DCs were capable of producing cytokines at frequencies similar to their CD4+ counterparts from age-matched healthy control blood, after stimulation with LPS and gamma-IFN (Supplementary Table 3).