Rational laboratory diagnostics of primary immunodeficiency disorders

Screening evaluation

Important components of the laboratory immunological diagnostics are basic screening methods (differential blood counts, total serum immunoglobulin levels – IgM, IgG, IgA, and IgE) and immunophenotypic analysis of peripheral blood cells to prove the integrity of the immune system as well as in vitro functional assays and specific immunizations, e.g. with recall protein (tetanus toxoid) or polysaccharide antigens (pneumococcal polysaccharides) or neo-antigens (tick-borne encephalitis) to test the functionality of the immune system [13].

According to the AWMF guidelines for diagnostics of PIDs, leukopenia, lymphopenia, neutropenia, or thrombocytopenia, revealed during basic screening, can be the first indication for PID and require, if persistent and associated with disease, further workup. Low numbers of cells might point to either decreased generation of cells, i.e. developmental block (e.g. in the case of SCID or CID, congenital neutropenia, Wiskott-Aldrich syndrome) or reduced survival of cells (e.g. in case of autoimmune neutropenia, hemophagocytosis). Additionally, secondary causes (toxic and viral effects or other losses) should be taken into account. Meanwhile, elevated numbers of cells such as eosinophilia in association with an increased susceptibility to infections or immune dysregulation might also be indicative of an underlying immune defect (e.g. in case of hyper-IgE syndrome, Omenn syndrome, immunodysregulation, polyendocrinopathy, enteropathy, X-linked [IPEX] syndrome) [11]. Finally, findings during morphological evaluation of a blood smear can also serve as a hint for PID (e.g. Howell-Jolly bodies in the case of congenital asplenia, microthrombocytes in the case of Wiskott-Aldrich syndrome, giant granular cells in the case of Chediak-Higashi syndrome) [11, 14].

According to the ESID registry, >50% of patients with PID apparently experience ADs; therefore, assessment of serum immunoglobulin levels is an important test in case of suspicion of PID. Elevated levels of immunoglobulins might also suggest an immunodeficiency, especially increased IgE levels (e.g. in the case of hyper-IgE syndrome, Omenn syndrome, IPEX syndrome), high levels of IgM (e.g. in the case of class-switch recombination [CSR] deficiency) or elevated IgG levels (e.g. in the case of ALPS or inflammatory bowel disease) [12]. Determination of serum levels of IgG subclasses has particular indications and should not be used as a primary screening assay [12, 14]. It is rather more important to assess specific antibodies. In particular, antibodies to tetanus toxoid and diphtheria toxoid are robust assays to measure antibody responses to recall protein (T-cell-dependent) antigens. Pneumococcal polysaccharide vaccine is recommended to test the antibody response against polysaccharide (T-cell-independent) antigens and when required – in the case of immunoglobulin substitution – testing of neo-antigens (tick-borne encephalitis) [12–14].

Generally, a basic eight-color flow cytometric panel, including the cell surface markers CD45, CD3, CD4, CD8, CD19, CD16, CD56, and HLA-DR, is sufficient to discriminate among major lymphocyte populations, T, B, and NK cells, as well as to measure the “activation status” of CD4⁺ helper T cells and CD8⁺ cytotoxic T cells. However, a deep immunophenotyping of lymphocyte subpopulations is required in some cases, and these tests can be performed only in specialized immunological laboratories. For several decades, there has been no standardized approach to immunophenotyping of peripheral blood cells, sometimes leading to confusing nomenclature. A recent study has developed a comprehensive and practical approach to standardize the immunophenotyping of most T-, B-, and NK-cell subpopulations, which permits differentiating whether abnormalities or developmental shifts observed in lymphocyte subpopulations originate either from primary or secondary immunological disturbances [15]. There is an ESID-authorized Internet-based open-accessible platform – “Immune Phenotyping in Immunodeficiency” (IPID) that serves scientists and clinicians in the field of phenotyping of immune cells in immunodeficiency (http://www.ipidnet.org/index.html). Moreover, it is worth to mention another European initiative – the EuroFlow project in the field of standardized diagnostic flow cytometric immunophenotyping [16].

Panhypogammaglobulinemia includes X-linked agammaglobulinemia (XLA), transient hypogammaglobulinemia of infancy, common variable immunodeficiency (CVID), autosomal recessive B-cell deficiency, and combined T- and B-cell immunodeficiency with hypogammaglobulinemia [9]. As a general rule, lack of B cells excludes transient hypogammaglobulinemia of infancy and other causes of primary hypogammaglobulinemia, such as X-linked hyper-IgM syndrome [17]. Absence of B cells supports the diagnosis of XLA in a male subject, but the clinical phenotype can be mimicked by rare mutations in genes encoding the μ heavy chain, Igα, Igβ, λ5, B-cell linker (BLNK), leucine-rich repeat-containing (LRRC8), or the p85α subunit of phosphoinositide 3-kinase (PI3K) [17–19]. Moreover, lack of B cells in the periphery might also be implicative of IKAROS deficiency [9].

Variably low numbers of B cells and low to absent memory B cell as well as switched memory B-cell compartments are reported for several PIDs, e.g. DOCK8, LRBA, or CD27 deficiency [9]. Therefore, an expanded flow cytometric panel for B-cell deficiencies should include at least the following additional six B-cell differentiation markers, CD19, IgM, IgD, CD21, CD27, and CD38, in order to define CD19⁺CD38^hiIgM^hi transitional, CD19⁺IgM⁺IgD⁺CD27^- naïve, CD19⁺CD21^lowCD38^low innate-like, CD19⁺IgM⁺IgD⁺CD27⁺ marginal zone-like, CD19⁺IgD^-CD27⁺ class-switched memory B-cells, and IgM^-CD38^hi class-switched plasmablasts in the periphery [20].

As, during the first months of life, serum IgGs are predominantly of maternal origin, normal IgG serum levels can be detected, even in patients with impaired ability to produce antibodies. Very low levels of serum IgG and IgA with normal to increased serum IgM levels are suggestive of CSR defects caused by either intrinsic B-cell problems (AID or UNG deficiency) or impaired cross-talk between T and B lymphocytes (CD40L, CD40 deficiency or NEMO defect) [9, 12].

Lymphopenia and low numbers of T cells are a hallmark of SCID [9]. However, a possible HIV infection must be excluded in all SCID cases by PCR, as serology can be false negative. Thereby, it is very important to compare lymphocyte counts with those of age-matched healthy control subjects [21]. The presence of maternal T-cell engraftment or of residual autologous T cells in patients with CID might result in relatively preserved and even normal T-cell counts; however, in these cases, most circulating T cells have an activated/memory (CD45R0⁺) phenotype and there is a severe reduction of naive (CD45RA⁺) T lymphocytes [12]. Detection of CD4⁺CD31⁺ recent thymic emigrant T cells is another additional helpful step to check whether there is thymic T-cell output into the immunological periphery. During T-cell development, T-cell receptor excision circles (TRECs), consisting of circularized signal joints, are generated as a byproduct of V(D)J recombination and are exported to the periphery by newly generated T cells that leave the thymus. Levels of TRECs in circulating lymphocytes can be measured by means of PCR. TRECs are particularly high in newborns and infants (reflecting active thymic function), whereas in SCID patients, they are severely reduced [22, 23].

Markedly decreased numbers of T cells and significantly low counts of B cells in the periphery are indicative of T^-B^- SCID, including RAG1/2, ARTEMIS, PRKDC, AK2, and ADA deficiencies.

Disease-associated proteins can also be evaluated by flow cytometry to define the diagnosis. Regulatory T cells (T_Regs) are usually missing in IPEX. T_Regs are defined as CD4⁺CD25^brigh T cells that express intracellular FOXP3. FOXP3 is not functional in patients with IPEX, and only a residual expression is detected in some of IPEX patients [24].

Diagnosis of leukocyte adhesion deficiency type I (LAD1) is based on flow cytometric assessment of CD18 expression on the surface of leukocytes. Partial defects (2%–10% of normal density of CD18 molecules at the cell surface) are associated with a moderate form of the disease that permits more prolonged survival [25].

Protein-specific flow cytometry assays can also be used in the diagnosis of CSR deficiencies – in case of hypogammaglobulinemia, PCJ pneumonia, and neutropenia (CD40 expression on B cells) [26], MHC II defect – in case of normal CD8+ T-cell counts, but CD4+ T-cell lymphopenia (absence of HLA-DR expression) [27], WAS – in case of (micro-)thrombocytopenia and eczema (lack of WAS protein) [28], SCIDX1 – in case of SCID in male (absence of common γ chain) [29], XLA – in case of hypogammaglobulinemia and severe B-cell defect in male (lack of BTK protein in B cells and significantly decreased expression levels in monocytes) [30], DOCK8 – in case of high levels of IgE, food allergy, eczema, viral infections, and T-cell lymphopenia (expression of DOCK8 protein in lymphocytes) [31], ZAP-70 – in case of CID, severe CD8+ T-cell lymphopenia with normal CD4+ T-cell counts (expression of ZAP-70 protein in lymphocytes) [32], XLP1/2 – in case of EBV infections in male (expression of SAP and XIAP proteins in lymphocytes, respectively), and HLH caused by perforin deficiency – in case of clinical symptoms according to HLH criteria (expression of perforin in NK and T cells) [33].

However, interpretation of these assays should take into account that some mutations are permissive for residual protein expression [12] and that only a negative result is valid.

Advanced immunological and functional assays

Functional testing of immune cells, including in vitro proliferation and differentiation, cytokine production, phosphorylation of signaling proteins, induction of certain activation markers, and analysis of cytotoxicity, is the next logical step in the laboratory-based diagnostic workup of a patient with suspected PID. A growing number of functional assays are being offered as standard tests by specialized immunological diagnostic laboratories. However, many functional tests are still performed with appropriate controls on research basis only.

Stimulations with phytohemagglutinin, anti-CD3 antibodies, and anti-CD3+anti-CD28 antibodies (TCR co-stimulation) in vitro can be used to assess in vitro T-cell responsiveness to mitogens and TCR-dependent signaling. In patients with SCID, in vitro T-cell responses to mitogens is completely absent, whereas the numbers and proliferative responses of circulating T cells are often variable in patients with different forms of CIDs [12]. In case of impaired in vitro proliferation upon TCR-dependent signaling (anti-CD3 and anti-CD3+anti-CD28 stimulations), it is reasonable to test an activation capacity of T cells by measuring early activation markers (e.g. CD69, CD25) during stimulations in vitro. Analysis of cytokine production upon in vitro stimulations are performed to assess effector functions of T-cell subsets, namely Th1 (e.g. IFNγ, TNFα), Th2 (e.g. IL4, IL10), and Th17 (IL17) cells in the periphery. Induction of CD40L can be analyzed upon in vitro stimulation to prove or disprove the suspicion of a CD40L deficiency. Evaluating the “phosphorylation status” of signaling proteins (e.g. phospho-flow assays for pSTATs) within the first 15–30 min during specific stimulations will help to reveal defects in the intracellular signaling cascade of immune cells. Additionally, TCR repertoire can serve as a valuable diagnostic tool for differential diagnostics of leaky SCID and in the case of Omenn syndrome [34].

Investigation of patients with putative TLR-signaling defects can be facilitated using a screening assay to measure IL6 and TNFα production on stimulation of whole blood with TLR agonists, or by the failure of affected neutrophils to shed CD62 ligand following stimulation in vitro [12, 35].

Diagnosis of chronic granulomatous disease (CGD) is most commonly based on evaluation of dihydrorhodamine 123 (DHR-123) oxidation, as assessed by flow cytometry [36]. This test also permits identification of carriers of X-linked CGD, who have two populations of neutrophils, only one of which is capable of mediating DHR-123 oxidation [12, 36]. Patients with autosomal recessive CGD often have very modest but detectable levels of activity, as detected by this assay.

Other clinically relevant functional assays include the demonstration of markedly reduced NK cytotoxicity (measured against K562 erythroleukemic target cells) in patients with familial forms of HLH and impaired Fas-mediated apoptosis in patients with autoimmune lymphoproliferative syndrome (ALPS) [33, 37].

Measurement of hemolytic activity of the classical (CH50) and alternative (AP50) pathways of complement and subsequently of single complement components might guide in the diagnosis of complement deficiencies [38].

Whether or not the findings of laboratory diagnostics could prove the suspicion of PID, genetic diagnostics is the final, and decisive step in PID diagnostics, providing detailed molecular genetic insights into the pathomechanism of an underlying PID.

Genotype-phenotype correlation

The current classification of PIDs comprises more than 300 different diseases [9]. For the major part, the molecular cause and underlying gene or genes have been identified and can be analyzed to detect disease-causing mutations (Table 1). PID-causing genes affect nearly all components that function in the immune system and are involved in maturation of hematopoetic cell lineages, signaling, class switch, V(D)J recombination, tolerance induction, defense against specific pathogens, and also in pathways for which it is not immediately evident how they relate to immunodeficiency, e.g. mutations in a catalytic subunit of the glucose-6-phosphatase result in severe congenital neutropenia [39].

Today, the most common approach to diagnosing PIDs employs first the clinical examination of the patient, including family history, followed by the functional characterization of immunological components [10]. Finally, guided by the patient’s clinical and immunological findings, one or more candidate genes can be analyzed to give a definite molecular diagnosis. Unfortunately, determining which genes to assess is often complex due to several reasons.

1. Up to date, more than 200 genes are known to cause PIDs, and the number of newly discovered genes is increasing rapidly since the employment and advances of new technologies, i.e. whole exome and genome sequencing (WES and WGS, respectively) by next-generation sequencing (NGS) [40]. Compared to the previous version of 2011, 30 novel gene defects are reported in the current catalog of the IUIS [9]. Recently, Itan and Casanova [41] expanded in silico the list of potential PID-causing genes to 3110 based on genes predicted by the human genes connectome to be biologically close to known PID genes.

2. The clinical phenotype within a genotype can vary significantly. Different mutations in the same gene may result in completely different clinical and immune phenotypes, due to different effects on the protein function (F. Hauck and C. Klein, unpublished observations). For example, null mutations in RAG1 or RAG2 genes cause T-B-NK+SCID, whereas hypomorphic mutations in the same genes can associate with Omenn syndrome, leaky SCID, or CID with granulomas [42]. Even the same mutation in the same gene can lead to variable clinical pictures in different patients due to additional influences such as modifier genes, environmental factors, or epigenetic modifications.

3. Similar clinical phenotypes can be caused by more than one genotype. For example, six genes are known to date to be associated with T-B+NK+ SCID [9].

4. There are conditions – reflected in the newly added category “phenocopies of PID” of the IUIS classification – that resemble inherited immunodeficiencies but are not due to germline mutations, instead arising from acquired mechanisms such as somatic mutations in TNFRSF6 associated with the ALPS (ALPS-sFAS) [9].

5. About 30% of patients who are referred to an immunodeficiency clinic have a distinct immunodeficiency but present with a clinical and cellular phenotype that is not well described in the literature [43]. Furthermore, for most patients with CVID, which comprises about 55% of PID patients based on ESID statistics, no genetic defect has been identified [44].

However, molecular characterization of the underlying immune defect is important and has several implications for the medical management of the patients. With the knowledge of the genetic cause, an early and specific therapy and the appropriate genetic counseling is possible. In some cases, the genotype may help to anticipate the course of the disease. A strong genotype-phenotype correlation exists for example for the WAS gene. Mutations predicted to abrogate protein expression are typically associated with the classical Wiskott-Aldrich syndrome, whereas missense mutations, resulting in the expression of a mutated protein, typically cause X-linked thrombocytopenia, the milder form of the disease [45]. However, unexpected mutations might be discovered in patients even with milder phenotypes or highly variable clinical courses. Hence, decisions on the therapy should never be derived from the genotype alone. Identification of the genetic cause further allows prenatal diagnosis and carrier identification and therefore permits presymptomatic identification of patients affected with life-threatening forms of PIDs, allowing early interventions. In atypical presentations, mutation analysis can help to establish a diagnosis. Beside the clinical implications, the characterization of the molecular defect provides important knowledge base about the pathophysiology of the disease and the immune system in general.

Current genetic approaches

Different methods are available for the molecular genetic analysis of PIDs: for a long time, Sanger sequencing was the only choice and the gold standard to identify causal mutations, but NGS techniques are becoming increasingly accessible in clinical laboratories, enabling new diagnostic possibilities.

For the analysis of single genes in distinct PID entities or if the results of previously performed tests allow to narrow the list down to only a few possible candidate genes, Sanger sequencing is and will most likely remain the method of choice. If more genes are included in the candidate list or previous results do not give any clear hints, Sanger sequencing becomes too expensive, time-consuming, and inefficient. Laboratory time and effort will be enormous compared to the high-throughput approach of NGS, where hundreds of genes or even can be sequenced (WES and WGS, respectively) all at once. NGS has already been widely used in research facilities to identify novel disease-causing genes, including PID genes [46]. Recently, NGS has been proved to be a sensitive and accurate method feasible for the routine molecular diagnosis of PIDs in the clinical setting as well [47–49].

All NGS technologies are based on the massively parallel sequencing of millions of DNA fragments, resulting in very high numbers of sequenced bases per run. Sequencing of different target regions in parallel is possible and dependent on the size of target region many samples can be multiplexed in one sequencing run due to sequence barcodes. The general NGS workflow encompasses the preparation of templates from genomic DNA, sequencing, data processing and analysis, and finally filtering of identified variants and their interpretation. For each step, several possible techniques or software solutions are available, provided by different vendors, all having their specific advantages and limits that have to be considered [50, 51]. Therefore, implementing NGS for clinical diagnostic purposes is complex and essentially requires great care in regard to quality assurance of each step in the complete process [52, 53].

Three sequencing approaches can be applied for mutation search depending on the size of the desired target region to be analyzed: (1) targeted resequencing of known PID-related genes, (2) WES, which includes all protein-coding exons, harboring approximately 85% of deleterious sequence variants [54], and (3) WGS covering the complete genome, in addition comprising introns, regulatory elements, and promoter sequences.

A stepwise process for obtaining a molecular diagnosis is likely the most efficient way in regard to costs and turnaround time. The first step would be searching for the causal gene by targeted resequencing of a panel of multiple genes, containing either all known PID genes or alternatively a smaller subset of genes, which are associated with a certain phenotype, e.g. genes related to syndromes presenting with autoimmunity, neutropenia, or CID.

The relatively small target region size of a gene panel allows analyzing several samples in parallel, which reduces costs and generally results in a higher sequence coverage compared with WES or WGS. Interpretation of the restricted data output of genes that are already better characterized with respect to its common variants and mutations is more convenient than to deal with the enormous amount of data generated by WES and WGS. However, NGS panels have to be updated regularly to include newly identified PID-associated genes and the diagnostic yield of the targeted approach remains to be evaluated. Nijman et al. [47] reported only four new genetic diagnosis in 26 patients, although the sensitivity to detect point mutations was higher than 99.9%. If no causative variants are found in the analyzed gene regions, the investigation can be expanded to WES or even WGS to search for mutations in yet unidentified PID-related genes including non-coding regions, which are often important for regulation of coding genes [40, 55]. In an exome analysis, an average of more than 20,000 single-nucleotide variants are identified, which are subsequently filtered, e.g. with respect to frequency and mode of inheritance [56]. At the end, about 20–40 variants will remain for detailed assessment. Candidate genes are then further filtered based on functional relevance and expression. Bioinformatics software algorithms can be used to evaluate sequence conservation and predicted pathogenicity of the variants in silico [57]. However, a definite proof for the clinical relevance of a previously unknown sequence variant is only possible by the application of additional functional assays that cannot be provided by most of the routine diagnostic laboratories. Further limitations of the current NGS methods have to be kept in mind. The sensitivity for detecting sequence variations other than single nucleotide exchanges, i.e. complex structural variations such as large deletions, insertions, or inversions is still low [58]. Covering repetitive areas and genes with pseudogenes or highly homologous genes might be complicated, leaving sequence gaps that may have to be complemented by Sanger sequencing. Additionally, somatic mosaics as described, e.g. for cryopyrin-associated syndromes or ALPS-sFAS are likely not discovered by NGS, depending on the achieved sequence coverage. These restrictions should be considered in the genetic report in a way that it is understandable for physicians who might not have to interpret such reports every day.

Although there are limitations at present, NGS approaches have proven their efficiency, accuracy, and applicability in the molecular diagnostics of PIDs [47–49]. Rapid progress in the NGS technology itself (e.g. longer reads) as well as in algorithms for data processing, mapping of sequence reads, and filtering of variants will certainly further improve quality of data output and thereby the sensitivity of the method.