Flow Based Enumeration of Plasmablasts in Peripheral Blood After Vaccination as a Novel Diagnostic Marker for Assessing Antibody Responses in Patients with Hypogammaglobulinaemia

Hypogammaglobulinaemic patients are often started on immunoglobulin substitution therapy before antibody production is adequately evaluated. In such a situation, it is difficult to segregate transferred from antigen-induced specific antibody. Therefore we characterized changes in B-cell subpopulations in hypogammaglobulinaemic patients, including plasmablasts, in peripheral blood by flow cytometry after in vivo antigen challenge. We investigated the specificity of antibody production on the B-cell level by ELISPOT, which is independent of substitution therapy.

patients (Cunningham-Rundles & Bodian 1999;Schaffer et al. 2007). CVID, therefore, is a heterogeneous group of patients expected to have multiple etiologies, all sharing similar immunologic and clinical characteristics.
Although the precise pathogenesis of CVID remains unknown, a number of common abnormalities involving peripheral blood lymphocytes were described including differences in the number of naïve B cells (follicular B cells), CD21 low B cells, transitional B cells, nonclass-switched IgM/IgD memory B cells (marginal zone-like B cells) (Klein et al. 1997;Shi et al. 2003;Tangye & Tarlinton 2009), class-switched memory B cells and plasmablasts (Carsetti et al. 2004;Sanchez-Ramon et al. 2008;Weller et al. 2004). Specifically, CVID patients have reduced populations of CD27 + memory B cells (classswitched memory B cells and marginal zone-like B cells) and increased percentages of undifferentiated B cells (immature CD21 low B cells (Rakhmanov et al. 2009) and naïve CD27 -B cells) associated with impaired class switching (Piqueras et al. 2003;Warnatz et al. 2002) and poor differentiation into plasma cells (Taubenheim et al. 2005) when compared to a control population (Ferry et al. 2005;Litzman et al. 2007).
In addition, a vast array of T-cell abnormalities has been described in CVID patients, including defects in TCR-dependent T-cell activation (Thon et al. 1997), reduced frequency of antigen-specific T cells, impaired IL-2 release in CD4 + T cells (Funauchi et al. 1995), decreased lymphocyte proliferation to mitogens and antigens , lack of generation of antigen-primed T cells after prophylactic vaccination (Bryant et al. 1990;Giovannetti et al. 2007), impaired cytokine production ; Thon et al. 1997), reduced expression of CD40L on activated T cells (Farrington et al. 1994;Piqueras et al. 2003; Thon et al. 1997;Warnatz et al. 2002), significant decrease in Treg cells in CVID patients with granulomatous manifestations and immune cytopenias (Horn et al. 2009), significant reduction of frequency and absolute counts of CD4 + T cells, percentage increase in CD8 + T cells, decrease in distribution of CD4 + and CD8 + naïve T cells in comparison to healthy controls (Giovannetti et al. 2007;Mouillot et al. 2010). This complex list of T-cell abnormality likely plays a major role in determining the clinical course of CVID patients.
In spite all of these multiple T-cell defects proposed as possible cause of CVID, the classification schemes presently in use are based on functional or phenotypic characteristics of B cells (assessment of immunoglobulin synthesis in vitro and phenotypic subsets of peripheral blood B cells): Bryant British classification (Bryant et al. 1990), Freiburg classification (Warnatz et al. 2002), Paris classification (Piqueras et al. 2003) and the recent EUROclass classification (Wehr et al. 2008). A few authors, however, suggested T-cell phenotyping as an aditional parameter for classifying CVID, and current efforts aim at the definition of combined T and B-cell phenotyping for the classification of CVID (Mouillot et al. 2010;.
Although a lot is known about B cell subsets in of CVID patients, the way their B-cell subpopulations change in response to vaccination compared to normal individuals is largely unknown. Specifically, there are limited data as to antibody responses to protein or polysaccharide antigens and the quantity and quality of antibodies produced by patient from different groups of CVID patients.
We focused on (1) specific in vitro antibody production by individual B cells following vaccinations by T-dependent (protein) and T-independent (polysaccharide) antigens and (2) www.intechopen.com changes of B-cell subpopulation after vaccination in peripheral blood of CVID patients and healthy donors (Chovancova et al. 2011).
The red-coloured spots were counted with the AID ELISPOT reader (AID, Autoimmun Diagnostika GmbH, Strassberg, Germany). This provided accurate recognition and calculation www.intechopen.com of the spots and allowed objective differentiation between background and "real" spots. The results were expressed as a number of SFC per million B cells.

Immunization of subjects
Thirty-seven patients with established CVID (14 males, 23 females, age range 20 -74 years) were examined. Twenty-six patients were treated with regular infusions of intravenous immunoglobulin (IVIG), six patients received regular subcutaneous immunoglobulin (SCIG) injections and one patient intramuscular immunoglobulin therapy (IMIG). Four patients were newly diagnosed and not yet on immunoglobulin replacement therapy at the time of the study.
All CVID patients were vaccinated simultaneously with tetanus toxoid (TET) vaccine (ALTEANA, Sevapharma, Prague, Czech Republic) and unconjugated pneumococcal polysaccharide (PPS) antigens (PNEUMO 23,Sanofi Pasteur,Lyon,France), except patient no. 34, who received PPS one year after TET. All patients on IVIG were vaccinated one week prior to administration of replacement therapy.
The control group consisted of 80 healthy individuals. Fifty (16 males, 34 females, age range 22 -72 years) were vaccinated with TET; ten (4 males, 6 females, age range 15 -46 years) were given PPS alone; twenty (8 males, 12 females, age range 14 -50 years) received both TET and PPS. The study was approved by the Ethics Committee of Masaryk University, Brno and signed informed consent was obtained from each participant.

Enzyme-linked immunosorbent assay (ELISA) and immunoglobulin quantification
Commercially available kits were used for measuring specific IgG antibody levels against tetanus toxoid (VaccZyme TM Human Anti Tetanus Toxoid IgG EIA Kit, The Binding Site Group Ltd, Birmingham, United Kingdom) and IgG antibodies titers against IgA (Human Anti-IgA isotype IgG ELISA, BioVendor, Brno, Czech Republic) in serum.
Trough serum levels of immunoglobulins IgG, IgA and IgM were measured in CVID patients prior to the IVIG infusion by nephelometry using the BN2 Nephelometer (Dade Behring, Marburg, Germany) according to the manufacturer's instructions.

Statistical analysis
Data were analyzed using the STATISTICA software [StatSoft, Inc. (2007), STATISTICA (data analysis software system), version 8.0.; www.statsoft.com]. Mann-Whitney U-test and Wilcoxon matched pairs test were used for analyses of dependencies between particular parameters in studied groups; p < 0.05 was regarded as statistically significant.

Kinetics and optimal timing for detection of specific spot forming cells isolated from peripheral blood after vaccination
The kinetics of anti-TET (T-dependent) specific antibody production by peripheral blood B cells was tested by ELISPOT assay in healthy volunteers from day 5 to day 9 after antigenic challenge. The same strategy was used in the assessment of anti-PPS (T-independent) specific antibody production in healthy controls from day 1 to day 8 after antigen challenge. Day 7 was found to be optimal for the detection of specific antibody producing B-cells in peripheral blood for both antigens and all tested immunoglobulin isotypes (IgG, IgA, and IgM). Our findings are in agreement with previous studies (Kodo et al. 1984;Stevens et al. 1979;Thiele et al. 1982).

Kinetics and specific antibody responses against protein (T-dependent) and polysaccharide (T-independent) antigens in healthy individuals
The group of healthy controls was vaccinated with protein antigen (tetanus toxoid, TET), unconjugated PPS antigens (PNEUMO 23) either separately or in combination. We found no significant difference in the number of SFC (IgG, IgA, IgM) against vaccinated antigens whether they were administered separately or simultaneously (Mann-Whitney U-test, p with range between 0.56 to 0.98). The number of specific SFC against both types of vaccines in the cohort of healthy controls is shown in Table 1

Specific antibody response in subgroups of CVID patients
CVID patients (n = 37) were classified according to the Freiburg (Warnatz et al. 2002) and EUROclass classification (Wehr et al. 2008) (Table 2), allowing a comparative analysis of antibody production and clinical phenotype. As we had expected, the majority of our welldefined CVID patients did not mount a specific humoral immune response against the two vaccines but several patients produced low numbers of vaccine-specific SFC (see below).
As for EUROclass classification scheme, 3 patients of group smB+21 norm (n = 7, patient no. 18, 19, 20), 1 patient of group smB+21 low (n = 6, patient no. 13) and 1 patient of group smB-21 low (n = 12, no. 1) had detectable IgG antibody responses against tetanus toxoid. In group smB+21 low there was 1 patient (no. 14) who secreted IgM and another patient (no. 12) who formed IgA and IgM antibodies against PPS. The latter patient is the only one among the CVID group who formed specific antibodies of 2 different immunoglobulin isotypes.
Regarding the group smB-21 norm (n = 12), no specific antibody production was detected. In the Freiburg classification all patients with detectable antibody responses (no. 12, 13, 14, 18, 19 and 20) were from group II the exception (no. 1) being a group Ia patient.   The decreased production of SFC in CVID patients was independent of replacement immunoglobulin treatment: four CVID patients without substitution therapy showed the same defect in the production of SFC and specific antibodies after vaccination as CVID patients on replacement therapy.

Changes of B-cell subpopulations in peripheral blood one week after vaccination
The mean percentage of CD19 + B cells was 11 ± 4 % in healthy controls and 13 ± 7.6 % in CVID patients before vaccination. One week after vaccination the percentages were unchanged (12 ± 5 % in healthy controls and 13 ± 6.7 % in CVID patients).
We then examined the changes of absolute and relative numbers of plasmablasts and other B lymphocyte subpopulations in the peripheral blood one week after antigen challenge (Fig.  1, 2). In healthy controls no statistically significant changes in absolute and relative numbers of switched memory B cells were found between the two measurement time points, before www.intechopen.com  and one week after vaccination. However, a highly significant increase in absolute as well as relative numbers of plasmablasts gated as IgD -CD27 ++ (PB CD27 ++ ) cells and IgM -CD38 ++ (PB CD38 ++ ) cells (p<0.001 in both cases) occurred (Fig. 3), while the absolute and relative numbers of CD21 low B cells (p<0.02), naïve B cells (p<0.001) and MZ-like B cells (p<0.001) decreased. In contrast, among the cohort of CVID patients no statistically significant changes of examined cellular subpopulations, including plasmablasts (Fig. 1, 2 and 3) were observed except for a slight increase in smB cells to a level still well below the levels of healthy controls. This increase was statistically significant in Wilcoxon matched pairs test.
The fact that the number of plasmablasts corresponds with the number of SFC strongly suggest that the examination of peripheral blood plasmablasts on day 7 after vaccination can be used as a surrogate marker for specific antibody responses in normal controls and as a diagnostic procedure to identified CVID and other patients with defect in terminal B-cell differentiation (Chovancova et al. 2011).

Hypogammaglobulinaemic patients and diagnostic vaccination
Poor vaccination responses to protein and polysaccharide antigens is essential for definitionbased diagnosis of CVID (Conley et al. 1999). Quantitative assessment of specific antibody in serum is routinely performed by ELISA assay. However CVID patients are often started on immunoglobulin substitution therapy before antibody production is adequately evaluated. In such a situation, it is difficult to segregate transferred from antigen-induced specific antibody. We have designed a in vitro functional measurement of antibody production on the B-cell level using the ELISPOT technique, which is independent of substitution therapy (Chovancova et al. 2011). In addition, we monitored changes in B-cell subpopulations, including plasmablasts, in peripheral blood by flow cytometry after in vivo antigenic challenge.
The defect in the antibody production and SFC reduction observed in a cohort of CVID patients are not secondary to Ig substitution since the same defects were also seen in four CVID patients before starting Ig replacement therapy. IVIG treated CVID patients were vaccinated exactly one week before administration of immunoglobulin substitution. In this manner the theoretically possible influence of immunoglobulin replacement therapy on the generation of SFC was reduced.
Prior to this study, specific antibody production in substituted CVID patients following vaccination had been evaluated in serum. Goldacker et al. (Goldacker et al. 2007) measured specific antibodies in serum by ELISA assay. The contribution of parallel immunoglobulin substitution on antibody titers was difficult to correct and required a relatively complicated vaccination formula. Based on these calculations the authors reported a decrease in serum antibody levels against T-dependent and T-independent antigens in CVID patients between IVIG infusions. Using a meningococcal polysaccharide vaccine, Rezaei et al. described decreased vaccination response against meningococcal polysaccharide measured in serum of CVID patients while on IVIG Rezaei et al. 2010). Immunization with a protein neoantigen, e.g. bacteriophage, and investigation of immune response with neutralization assay brought similar results (Ochs et al. 1971). Nevertheless, there is very little quantitative data correlating individual vaccination responses to proposed functional www.intechopen.com  www.intechopen.com Fig. 3. Development of plasmablasts after vaccination. Plasmablasts (red arrows) were gated from CD19 + B cells (gate in column 1) as IgD -CD27 ++ (column 2) and IgM -CD38 ++ (column 3). The cells were investigated before (day 0) and on day 7 after vaccination. HC -healthy control; CVID -CVID patient, PB27 ++ and PB38 ++ -plasmablasts classifications of CVID (Goldacker et al. 2007;). Our group of CVID patients was arranged according to the Freiburg (Warnatz et al. 2002) and EUROclass classification (Wehr et al. 2008). As expected, the majority of our well-defined CVID patients (30/37) failed to mount a specific humoral immune response w h e n a n a l y s e d b y S F C s c o l l e c t e d f r o m peripheral blood before and after immunization. The seven CVID patients who responded had much smaller quantities of specific SFC compared to healthy donors. All but one patient with measurable antibody responses belong to group II of the Freiburg classification or EUROclass group smB + which represent those CVID patients with nearly normal numbers of classswitched memory B cells. Patients in these groups are characterized by milder complications of the disease compared to other groups (Alachkar et al. 2006;Wehr et al. 2008).

Novel diagnostic tool using flow cytometry in hypogammaglobulinaemic patients with vaccination
During the last few years a number of studies described differences between B-cell subpopulations of CVID patients and those of healthy volunteers but the kinetics of these changes after encounter with an antigen in vivo (Pinna et al. 2009) has not previously been explored. We investigated the dynamic changes of CD21 low B cells, naïve B cells, marginal zone-like B cells, plasmablasts and switched memory B cells of CVID patients compared to healthy donors (Chovancova et al. 2011). Previous studies showed that memory B cells and plasmablasts have different kinetics in peripheral blood (Stevens et al. 1979). Plasmablasts reach their peak on day 7 after encounter with the antigen in peripheral blood while switched memory B cells showed a marked increase in number on day 14 after antigen challenge (Pinna et al. 2009). The absolute number of naïve B lymphocytes is determined by the generation of new naïve B cells from the bone marrow pool (a slow process) and by acute loss of naïve B lymphocytes via further maturation after antigen encounter (Agenes et al. 2000). Statistically significant up-regulation of naïve B cells and its continued accumulation after antigen challenge in CVID patients indicates disturbed conversion of undifferentiated B cells to more mature B-cell stages in germinal centers. Differentiation is crucially dependent on T-lymphocyte help, suggesting that the basic defects in the majority of CVID patients are not in B cells but in helper T-lymphocytes (Borte et al. 2009;Fischer et al. 1996;Thon et al. 1997).
The reduced numbers of switched memory B cells which correlate with clinical complications (Ko et al. 2005;Viallard et al. 2006) and failure to increase the number of plasmablasts after antigen challenge may be explained by insufficient signals from helper T cells of CVID patients. In previous studies we and others have shown that B cells of CVID patients are able to produce antibodies if they are exposed in vitro to helper T-lymphocyte from healthy donors or to appropriate cytokines (Borte et al. 2009;Fischer et al. 1996;Thon et al. 1997) , 55 . Taubenheim et al. studied B-cell differentiation in lymph nodes from three CVID patients with splenomegaly and found distinct blocks in terminal plasma cell development but normal expression of a key regulator of terminal plasma cell differentiation, Blimp-1 (Taubenheim et al. 2005). Moreover, the clinically important observation that B cells from CVID patients may produce antibodies under certain circumstances correlates with the fact that CVID patients lacking IgA are able to generate IgG anti-IgA antibodies in vivo (Horn et al. 2007). Among our cohort of vaccinated CVID patients, two patients from subgroups Ib and II (Table 2) produced IgG anti-IgA antibodies in low titers although these two patients did not respond to vaccination.
Our observation that the majority of CVID patients lack antigen specific spot forming B cells and fail to increase circulating plasmablasts following in vivo antigen challenge provides a rapid screening test to demonstrate defective antibody responses in CVID patients, even when on replacement IVIG therapy (Chovancova et al. 2011).

Conclusion and clinical implications
Identification of circulating plasmablasts after vaccination is a new simple flow based test to assess antibody responses in hypogammaglobulinaemic patients, even if on immunoglobulin (IVIG or SCIG) replacement therapy.