Blood and serum collection

This sampling schedule applies to participants undergoing immunological evaluation for suspected humoral immune dysfunction, rather than routine vaccination recipients.

• 1.

  On the day of vaccination (baseline visit), collect 3 to 5 mL of peripheral blood from each participant into two collection tubes: a plain tube for serum collection and an EDTA tube for flow cytometry analysis.

• 2.

  Administer the Pneumovax23 PCP vaccine and the tetanus toxoid vaccine in accordance with institutional immunization guidelines and standard clinical practice.

• 3.

  Process the EDTA blood sample immediately following collection for whole blood memory B cell phenotyping by flow cytometry. Samples should be maintained at room temperature and protected from light prior to processing to preserve cell integrity.

• 4.

  Centrifuge the plain blood tube at 3,000 rpm for 5 min within 2 hours of collection to obtain serum. Aliquot a minimum of 1 mL of serum into appropriately labelled microcentrifuge tubes for enzyme immunoassay analysis.

• 5.

  A follow-up visit was conducted 28 to 30 days after vaccination to collect a second peripheral blood sample using a plain tube for post-vaccination serum analysis in all vaccine recipients.

• 6.

  Store all pre- and post-vaccination serum aliquots at −20°C until batch analysis is performed. Repeated freeze-thaw cycles should be avoided to maintain antibody stability.

Memory B cell phenotyping (whole blood lyse-wash method)

Flow cytometry acquisition, quality control (QC), and gating strategy

• 1.

  Prior to sample acquisition, perform routine instrument QC and fluidics cleaning using FACS Flow™ and FACS Clean™ solutions in accordance with the manufacturer’s recommendations. Daily cytometer performance was verified using calibration beads to ensure consistent laser alignment, fluorescence sensitivity, and signal stability.

• 2.

  Compensation settings were established using single-stained compensation controls for each fluorochrome to correct for spectral overlap. Fluorescence-minus-one (FMO) controls were applied where appropriate to support accurate gating of dim populations.

• 3.

  A minimum of 50,000 lymphocyte events per sample were acquired to ensure sufficient resolution of B cell subsets.

• 4.

  Figure 1 demonstrates a gating strategy using the FlowJo software. Lymphocytes are first identified based on forward scatter (FSC) and side scatter (SSC) characteristics. Total B cells were defined as CD19⁺ events within the lymphocyte gate. Naïve and memory B cell compartments are initially distinguished based on IgD and CD27 expression. Further subclassification is performed using IgM, CD27, CD38, and CD24 markers to identify:

  • Naïve B cells (CD19⁺CD27⁻IgD⁺)

  • Transitional B cells (CD19⁺CD27⁻CD24⁺⁺CD38⁺⁺)

  • Total memory B cells (CD19⁺CD27⁺)

  • Non-switched memory B cells (CD19⁺CD27⁺IgD⁺IgM⁺)

  • Class-switched memory B cells (CD19⁺CD27⁺IgD⁻IgM⁻)

  • Plasmablasts (CD19⁺CD27⁺CD38⁺⁺)

• 5.

  Each cell captured will be analyzed according to the gated populations.

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Figure 1. Representative flow cytometry gating strategy for B cell subsets and memory B cell phenotyping.

ELISA for specific antibody response measurement

Bring VaccZyme™ ELISA kit and all reagents to room temperature (20–25°C) approximately 60 min before use. The 96-well ELISA plates are pre-coated with PCP or tetanus toxoid antigens according to the manufacturer’s specifications. All reagents should be gently mixed before use.

Serum samples, calibrators, and manufacturer-provided high and low controls are run in duplicate. Serum samples are diluted 1:100 using the supplied diluent (10 µL serum + 1,000 µL diluent). The wash buffer is prepared by diluting the concentrated wash solution 1:20 with distilled water.

A total of 100 µL of calibrators, controls, and diluted samples were added to the designated wells and incubated for 30 min at room temperature. Plates are washed three times with 250–350 µL wash buffer per well using an automated plate washer. Subsequently, 100 µL of conjugate solution is added to each well and incubated for 30 min at room temperature, followed by three additional wash cycles. Then, 100 µL of tetramethylbenzidine (TMB) substrate was added and incubated for 30 min at room temperature in the dark. The reaction is stopped by adding 100 µL of stop solution (3 M phosphoric acid), and optical density (OD) is read at 450 nm using a microplate reader within 30 min.

Data analysis and QC

Strict QC criteria are applied to every assay run. Calibrators and manufacturer-provided high and low controls are included in every batch run. Measured values for the high and low controls of anti-PCP or anti-tetanus IgG must fall within the acceptable ranges specified in the QC certificate supplied with each kit.

For each calibrator, sample, and control, the mean OD value of duplicate wells is calculated, and the coefficient of variation (CV) between duplicates is determined. Duplicate measurements are accepted only when the CV is < 15%; samples or controls exceeding this threshold are repeated.

Calibration curves for anti-PCP IgG and anti-tetanus toxoid IgG were generated using a four-parameter logistic (4PL) model in SkanIt™ microplate reader software (Thermo Fisher Scientific) to obtain the best fit, as illustrated in Figure 2. The standard curve must closely match the sigmoidal shape provided in the manufacturer’s QC certificate, and assay runs are accepted only when all calibrator points fall within the specified ranges, and the coefficient of determination (R²) is ≥ 0.98.

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Figure 2. Representative calibration curves for specific antibody responses are shown: (a) anti-PCP IgG antibody and (b) anti-tetanus toxoid IgG antibody. The four-parameter logistic (4PL) curve generated using SkanIt™ microplate reader software (Thermo Fisher Scientific) is displayed on the right, while the manufacturer’s reference curve is shown on the left. The similar curve shape between the two graphs demonstrates that a valid and reliable standard curve was achieved. PCP: pneumococcal polysaccharide. Adapted from Binding Site with permission. © 2026 Thermo Fisher Scientific Inc and its subsidiaries. All rights reserved. Unauthorized use, reproduction or distribution is prohibited.

Any calibrator or sample point deviating markedly from the fitted curve is treated as an outlier and excluded from analysis. Calibrator values are pre-adjusted for the 1:100 sample dilution; therefore, no additional dilution correction is required.

Antibody concentrations for anti-PCP and anti-tetanus IgG in samples and controls were calculated directly from the fitted calibration curve. Results are expressed as mg/L for pneumococcal antibodies and IU/mL for tetanus antibodies. Pre- and post-vaccination antibody concentrations are compared within subjects to determine the adequacy of antibody responses. Protective status is determined based on the post-vaccination antibody titre.

A protective pneumococcal antibody response is defined as an antibody concentration ≥ 110 µg/mL, representing an adequate overall pneumococcal IgG response, while a protective tetanus IgG response is defined as > 0.15 IU/mL. In this protocol, deficient pneumococcal or tetanus responses are indicated by a post-vaccination antibody concentration that fails to reach at least a fourfold increase compared with the pre-vaccination level.

Expected immunological and analytical

This protocol integrates direct whole blood staining for memory B cell phenotyping with ELISA-based quantification of vaccine-specific IgG responses, enabling simultaneous assessment of cellular and functional humoral immunity in children with recurrent infections or suspected antibody deficiencies. When performed according to the described QC parameters, the workflow is expected to generate biologically plausible B-cell subset distributions consistent with previously reported age-dependent distributions of B-cell subsets in healthy paediatric populations [21–24]. Interpretation of results requires age-stratified reference ranges, as B-cell subset composition varies throughout childhood. Population-specific reference intervals are recommended for clinical applications. Where external reference intervals are adopted, they should be verified in the local population to account for demographic and methodological differences.

Memory B cell phenotyping

Multiple studies have reported that naïve B cells typically comprise approximately 60–70% of circulating B cells in healthy children, while class-switched memory B cells account for approximately 10–20% and non-switched memory B cells for 10–15%, with plasmablasts generally representing less than 2% of total B cells under non-inflammatory conditions [21–24]. In children with recurrent infections or suspected humoral immunodeficiency, deviations from these distributions may be observed. Reduced proportions of class-switched memory B cells have been associated with impaired long-term immune memory and increased susceptibility to recurrent respiratory infections [14]. Similarly, increased proportions of transitional B cells may reflect delayed or defective B cell maturation, which has been described in patients with antibody deficiencies and IgG subclass deficiencies [7, 18]. These phenotypic patterns are consistent with previous reports demonstrating alterations in the memory B cell compartments in children with SAD and related disorders [7, 17].

It is important to acknowledge that the child’s age may influence B-cell subset composition and immune responsiveness. The paediatric immune system undergoes dynamic maturation, with younger children exhibiting higher proportions of naïve B cells and progressively increasing memory B cell compartments with advancing age [21–24]. This age-dependent variation is particularly relevant when interpreting the naïve and memory B cell populations. The present protocol does not specifically address age-related immunological differences but instead provides a standardized methodological approach applicable across paediatric age groups. Consequently, interpretation of B cell subset frequencies should be performed using age-matched reference ranges whenever available. Table 2 in this manuscript is intended as an example of how a laboratory can display the B cell subset results accordingly.

Vaccine-specific antibody quantification

ELISA-based measurement of PCP and tetanus toxoid-specific IgG provides a functional assessment of humoral immune competence. In immunologically healthy children, post-vaccination antibody concentrations are expected to reach protective thresholds and demonstrate a robust rise compared with baseline levels [8, 9, 25, 26]. Protective responses are typically defined as pneumococcal IgG concentrations ≥ 110 µg/mL and tetanus IgG concentrations > 0.15 IU/mL, and normal antibody responses by a minimum fourfold increase from pre-vaccination levels [9, 27, 28]. Table 3 represents how the outcome can be displayed in formal laboratory results. The antibody response findings must correlate clinically, especially in evaluating vaccine responsiveness in children with recurrent infections.

Children with SAD fail to achieve adequate protective antibody concentrations or demonstrate a sufficient post-vaccination increment despite normal total immunoglobulin levels [7–9, 27]. Such impaired vaccine responses have been consistently reported in paediatric cohorts with recurrent infections and are often accompanied by abnormalities in memory B cell subsets, particularly reduced IgM-memory and class-switched memory B cells [7, 17]. Therefore, the integration of memory B cell phenotyping with vaccine-specific antibody quantification in this protocol supports a more comprehensive assessment of humoral immunity and enhances interpretive confidence by linking cellular immune involvement with functional antibody production.

Overall, the workflow is intended to facilitate reproducible implementation, particularly for use in clinical and research laboratories establishing or expanding immunological assessment of children with recurrent infections.

Limitations

This protocol provides a standardized workflow for measuring memory B cells and vaccine-specific antibody responses in children with recurrent infections. However, several limitations should be acknowledged. First, this protocol is designed to provide supportive immunological insights and may help indicate the presence of an underlying humoral immune abnormality; however, it is not intended to serve as a stand-alone diagnostic tool for primary immunodeficiency. All results should therefore be interpreted in the context of clinical findings and routine immunological investigations, in accordance with established diagnostic guidelines. Second, the method requires fresh whole blood; delayed handling or transport may compromise B-cell viability and staining quality, and the protocol is not suitable for settings where PBMC cryopreservation is preferred. Third, flow cytometry results may vary depending on instrument configuration and fluorochrome compensation, necessitating local optimization. Finally, the manuscript does not include representative comparative data between healthy controls and patients with confirmed PID. While this limits demonstration of disease-specific patterns, the primary aim of the study is to present a standardized methodological framework rather than diagnostic validation. Future studies applying this protocol to well-characterized patient cohorts will be essential to establish disease-associated immunophenotypic patterns.

Discussion

This protocol presents a standardized workflow integrating whole blood memory B cell phenotyping with vaccine-specific antibody quantification to facilitate comprehensive evaluation of humoral immunity, especially among children with recurrent infections. By combining cellular immunophenotyping with functional serological assessment, the protocol provides complementary insights into both the structural integrity of the B cell compartment and the capacity to mount antigen-specific antibody responses. Such an integrated approach is particularly relevant in the clinical investigation of suspected PID, where isolated phenotypic or serological abnormalities may not fully capture immune competence, hence leading to missed/under interpretation of PID.

Flow cytometric analysis using the described gating strategy allows clear discrimination of major B cell subsets, including naïve B cells, transitional B cells, class-switched memory B cells, non-switched memory B cells, and plasmablasts. In healthy children, naïve B cells typically constitute the predominant population, reflecting ongoing B cell development, while memory B cells represent a smaller but stable fraction of circulating B cells [22, 29]. Memory B cell subsets, particularly class-switched memory B cells have been increasingly recognized as important indicators of long-term humoral immune competence and correlates of vaccine responsiveness. Several studies have demonstrated that children with SAD or IGGSD frequently exhibit diminished memory B cell compartments, particularly reduced class-switched memory B cells and IgM memory B cells, which have been consistently associated with impaired vaccine responsiveness and increased susceptibility to recurrent infections in paediatric populations [17, 30]. In common variable immunodeficiency (CVID) patients, reduced frequencies of CD27⁺ memory B cells are associated with impaired antigen-specific memory responses and suboptimal vaccine serology outcomes [31]. In this context, the combined assessment of B cell subset composition alongside post-vaccination antibody titres strengthens clinical interpretation by enabling differentiation between defects in B cell maturation and functional antibody production, enabling a more comprehensive interpretation of immune dysfunction than either approach alone.

PBMC isolation using Ficoll density gradient centrifugation remains a widely used method for immunophenotyping. While PBMC-based assays provide high-quality cellular preparations, they are labour-intensive, time-consuming, and prone to cell loss, particularly of fragile populations such as plasmablasts [32]. Delayed processing can further compromise cell viability and functionality, which is particularly challenging in paediatric settings. In contrast, whole blood lyse-wash staining offers several practical advantages. It minimizes sample handling, reduces processing time, and allows same-day flow cytometric acquisition [16]. Although the relative frequencies of most B cell subsets are generally preserved following PBMC isolation, the whole blood approach enhances operational efficiency and is more feasible in routine clinical laboratories, especially in resource-limited environments. This protocol therefore prioritizes practicality and reproducibility while maintaining reliable immunophenotypic resolution.

In future studies, several enhancements may further strengthen the utility of this protocol. Expansion of the antibody staining panel to include additional immunoglobulin subclasses, such as IgA1, IgA2, IgG1, IgG2, IgG3, IgG4, and IgE, would enable more detailed characterization of humoral immune defects, particularly in patients with subclass deficiencies. Furthermore, incorporation of antigen-specific memory B cell staining represents a promising direction for directly assessing vaccine-induced B cell memory. Antigen-specific probes are already available for tetanus toxoid and could potentially be extended to other clinically relevant vaccines, providing a direct cellular correlate of vaccine responsiveness. Such advancements would deepen mechanistic understanding of humoral immune dysfunction and further enhance the clinical relevance of the integrated workflow.

In summary, this protocol offers a practical and clinically meaningful framework for evaluating of memory B cell subsets and vaccine-specific antibody responses in children with recurrent infections. While not intended to replace established serotype-specific antibody assays, the protocol provides a feasible and reproducible framework that is particularly valuable in routine clinical immunology laboratories and resource-limited settings. In such settings, memory B cell subset profiling can serve as an important complementary marker of immune competence. The protocol also supports rapid turnaround time, facilitating timely clinical decision-making in paediatric patients presenting with recurrent infections. Importantly, the combined cellular and serological approach enhances diagnostic confidence. Future studies incorporating age-stratified reference data and patient cohorts with defined immunodeficiencies will further enhance its clinical applicability and diagnostic utility.