B cells grow up: Studies of B cell activation, proliferation and differentiation in primary antibody deficiencies - Thesis

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B cells grow up: Studies of B cell activation, proliferation and differentiation in

primary antibody deficiencies

aan de Kerk, D.J.

Publication date

2016

Document Version

Final published version

Link to publication

Citation for published version (APA):

aan de Kerk, D. J. (2016). B cells grow up: Studies of B cell activation, proliferation and

differentiation in primary antibody deficiencies.

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B cells

GROW UP

Daniel Jacob aan de Kerk

D

aniel Jac

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studies of B cell

activation,

proliferation

and differentiation

in primary

antibody deficiencies

B cells

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B CELLS GROW UP:

STUDIES OF B CELL ACTIVATION, PROLIFERATION

AND DIFFERENTIATION IN PRIMARY ANTIBODY DEFICIENCIES

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B cells grow up: Studies of B cell activation, proliferation and differentiation in primary antibody deficiencies Thesis, University of Amsterdam, The Netherlands

ISBN: 978-94-6182-706-7

Cover design, layout and printing: Off Page, Amsterdam

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B CELLS GROW UP:

STUDIES OF B CELL ACTIVATION, PROLIFERATION

AND DIFFERENTIATION IN PRIMARY ANTIBODY DEFICIENCIES

ACADEMISCH PROEFSCHRIFT

ter verkrijging van de graad van doctor aan de Universiteit van Amsterdam op gezag van de Rector Magnificus

prof. dr. ir. K.I.J. Maex

ten overstaan van een door het College voor Promoties ingestelde commissie, in het openbaar te verdedigen in de Agnietenkapel

op vrijdag 30 september 2016, te 10:00 uur

door

Daniel Jacob aan de Kerk geboren te Amsterdam

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PROMOTIECOMMISSIE

Promotor : Prof. Dr. T.W. Kuijpers Universiteit van Amsterdam

Copromotor : Dr. E.M.M. van Leeuwen Universiteit van Amsterdam

Overige leden: Prof. Dr. R.J.M. ten Berge Universiteit van Amsterdam

Dr. M.L. Boes Universitair Medisch Centrum Utrecht

Dr. M. van der Burg Erasmus Universiteit Rotterdam

Prof. Dr. T.B.H. Geitenbeek Universiteit van Amsterdam

Prof. Dr. S.M. van Ham Universiteit van Amsterdam

Prof. Dr. A.P. Kater Universiteit van Amsterdam

Prof. Dr. K. Warnatz Universiteit van Freiburg

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1 PEDIATRIC IMMUNE SYSTEM

Right from the start of life, newborns encounter a wide variety of pathogens. Various bacteria, viruses and parasites pose a threat to their health. Some motherly protection is provided by immunoglobulins (Ig) that crossed the placenta and new supply via breast milk. However this stock is limited, and not enough to keep protecting the child. In the course of evolution, humans have developed a very adequate immune system, which can protect the growing child against those pathogens. (1-4)

When immune defenses fail, clinical manifestations of a so-called immunodeficiency may develop. A distinction between primary and secondary immunodeficiencies is made to describe whether the defect is very likely or proven to be inherited but sometimes presents later during life, versus an acquired immunodeficiency because of another disease entity such as HIV infection, malignancy, immunosuppressive treatment. A primary immunodeficiency in fact exists already from birth onwards, but is manifests when the immune system is compromised due to interplay between genetic factors and the environmental triggers to which the host is being exposed. In some cases the exposures repeatedly take place and then finely undermine the host defense mechanisms permanently over time.

This thesis focuses on primary immunodeficiencies (PIDs), of which some are so mild they may go unnoticed for years and other types that are severe enough to be discovered almost as soon as an affected baby is born. (5)

ADAPTIVE IMMUNITY

The immune system is divided in two major categories. For direct and first line of defense we have the innate immune system, also known as the nonspecific immune system. Many cell types are involved, granulocytes (consisting of neutrophils, eosinophils, basophils), monocytes (which may develop into macrophages once infiltrated into the tissues), natural killer (NK) cells and the complement system, which consists of more than 35 different proteins circulating in plasma and a small series of complement regulatory proteins that are expressed on the surface of blood and tissue cells. The innate immune system provides an immediate response to pathogens. However, its components recognize only a limited number of pathogens (antigens) as pathogen-associated molecular patterns (PAMPs), expressed by foreign microbial species.

The second line of defense, called the adaptive immune system, is more complex creating an essential and critical immunological hallmark, i.e. long-lasting immunological memory. In contrast to the assumed presence of a transient form of innate memory, which is most likely determined by epigenetic imprinting on certain immune cells such as monocytes and such imprinted innate responses or trained immunity may be largely dictated by the half-life of the responding cell.(6)

In the case of adaptive immunity, after an initial or primary immune response to a specific pathogen or antigen, an enhanced secondary response will follow when encountering that same pathogen or antigen again. This specific immunological memory process is the basis of vaccination programs.

Major players of the adaptive immune system are B- and T-lymphocytes. B cells originate from the bone marrow whereas T cell precursors differentiate in the thymus gland. Each individual

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lymphocyte creates a unique antigen receptor on the B- and T cell, i.e. the B cell Receptor (BCR) and TCR, respectively, during their precursor differentiation. This creates a large repertoire of

different antigen receptors (around 1018_{combinations for T cells and 10}13_-1014_{for B cells), potentially}

recognizing many different antigenic determinants on a huge variety of pathogens. What makes them more unique is that, when the cell recognizes its specific antigen, it has the ability to proliferate in great numbers to rapidly generate enormous numbers of daughter cells that all will recognize the same target antigen. This expansion generates effector cells for long-term memory, so called B- and T-memory cells, and immunoglobulin-producing plasmablasts (PBs) and plasmacells (PCs). (7, 8)

NORMAL B CELL DEVELOPMENT

Normal B cell development starts in the bone marrow, where multipotent hematopoietic stem cells give rise to common lymphoid progenitors which can differentiate within the bone marrow niche into precursors B cells. Each B cell will have a unique B cell antigen receptor (BCR) a membrane-bound immunoglobulin that specifically binds antigen. Variation is created by a rearrangement process, called V(D)J recombination. A process in which one Variable, one Diversity and one Joining gene are randomly combined, ending up with numerous different functional BCRs. After completion of the BCR all B cells are tested to check for affinity for self-antigens, when this is the case those B cells will undergo apoptosis. All that ‘pass’ these developmental checks are ready to migrate to the periphery, at which stage they are called transitional B cells. (9, 10) The continuous stream of newly formed B cells leave the bone marrow, to circulate in the blood and pass through secondary lymphoid organs (lymph nodes, tonsils, spleen, Peyer’s patches and mucosa associated lymphoid tissue (MALT)) several times per day. There these antigen-naïve cells may get activated by antigens that have been captured by antigen-presenting cells (APCs), mostly dendritic cells (DCs), or by follicular dendritic cells (FDCs), specialized tissue cells that can capture the antigen from the continuous flow of lymph fluid percolating through the lymphoid tissues to keep and maximally expose certain antigens for a longer period of time.

Upon triggering by their specific antigen, naïve B cells become activated through the BCR, expand and differentiate into antigen-specific memory B cells or antibody-secreting PBs and PCs. Differentiation proceeds largely via two distinct pathways, depending on the nature of the stimulus, i.e. T cell-dependent antigens (typically protein structures) or a T cell-independent antigen (typically carbohydrates). (11, 12)

T cell-dependent differentiation leads to long-lived, highly specific, immunoglobulin-producing PCs. Activation results in the formation of germinal centers (GCs) in lymphoid organs

such as the lymph nodes, spleen and Peyer’s patches, where activated CD4+_{T cells interact with}

activated B cells through direct CD40L-CD40 interactions, and the release of soluble factors such as IL-21. These T-B cell interactions are essential for cellular processes that are needed to induce B cell differentiation into high-affinity, mostly IgG-producing, PCs. (13-16)

After specific antigen recognition, B cells have the unique ability to further optimize antigen-binding, the specificity, of their BCR. This is achieved by the introduction of point mutations in the V(D)J exons of their Ig heavy and light chains (somatic hypermutations; SHMs) and the subsequent selection for high-affinity mutants. Furthermore, the antibody effector functions

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can be modified by changing the isotype of the IGH constant region (Ig class-switch recombination; CSR). In this way, the initial IgM antibody may be exchanged for an IgG, IgA or IgE antibody. This process will increase the antibody’s functional avidity, complement-binding capacity and ability to be recognized by various immune effector cells such as the granulocytes, monocytes, macrophages or DC’s, by binding via the isotype-specific antibody tail (the so-called Fc domain) to the IgG, IgA or IgE receptors (FcγR, FcαR and FcεR, respectively). (17, 18)

T cell-independent activation most often leads to shorter-lived PBs producing specific antibodies of a more moderate affinity. In case of polysaccharide antigens this T-cell independent B cell activation is believed to occur without the elaborate GC formation and often does not result in a repertoire of high-affinity class-switched antibodies. When activated in a T cell-independent manner, B cells mostly differentiate into IgM-producing PBs, as demonstrated for the polyclonal activation either by BCR-independent Toll-Like Receptor (TLR) activation or BCR-dependent carbohydrate antigens through direct recognition of repetitive epitopes. (19-21)

PRIMARY ANTIBODY DEFICIENCY (PAD)

A defect in any of the stages of B cell development and differentiation can cause a primary antibody deficiency (PAD). PADs are the most common primary immunodeficiency in humans, characterized by a reduction or absence of serum immunoglobulins and poor antibody response to vaccinations. Over the past years the use of novel approaches, such as whole-exome or genome sequencing and mouse genetic engineering, have led to the identification of many genetic defects that cause PAD. Defects can be caused by B cell-intrinsic errors, or caused by functional impairments of other immune cells (e.g. T cells and innate immune cells), or by both. Studying these rare “experiments of nature”, i.e. natural mutants observed among the human PAD patients, has increased our understanding and characterization of the various molecular pathways involved in B cell development and antibody production. Among the B cell-intrinsic defects at least seven distinct groups can be recognized, namely defects in early B cell development, early survival, activation, T cell-dependent stimulation, T cell-independent stimulation, CSR and late survival (Figure 1). (5, 22-25)

PAD patients have increased susceptibility to infections, mainly bacterial that typically involve the upper and lower respiratory tract. Abscesses in the skin or other organs have also been described, as well as urinary tract infections and arthritis. Common infectious agents are encapsulated

Streptococcus pneumoniae and Haemophilus influenzae. (26-28) Early diagnosis and treatment

is important in PAD, next to optimal management of recurrent and chronic infections or serious complications such as autoimmunity, granulomatous inflammation, lymphoproliferation, and even malignancies that may occur. Starting immunoglobulin replacement therapy is therefore of great importance for all clinically affected PAD patients. Either subcutaneous of intravenous treatment regimens are available. These regimens, sometimes in combination with live-long antibiotic prophylaxis, may be necessary to control the number and severity of infections and the infection-related complications.

Even though the interest in PADs is growing and genetic techniques are improving fast, many of the PADs remain without a clear etiology. Genetic analysis of PADs proves to be more complex, new monogenetic causes are seldom found. Evidence suggests that different mutations in the same gene can even lead to distinct phenotypes. Also limited genotype–phenotype correlation have

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been observed thus far, suggesting that many additional genetic and as yet unidentified (and possible combinations of) environmental factors may contribute to the PAD pathogenesis. (25, 29)

DEFECTS IN EARLY B CELL DEVELOPMENT, CONGENITAL

AGAMMAGLOBULINEMIA’S

X-linked agammaglob ulinaemia (XLA) is a PAD associated with a complete defect in early B cell development. This condition is caused by mutations in the Bruton’s tyrosine kinase (BTK) gene and accounts for 85% of male patients diagnosed with agammaglobulinaemia. (30) Following pre-BCR engagement in early B cells (and BCR engagement in mature B cells), BTK is phosphoryl ated and is a key molecule in pre-B cell activation and differentia tion. Most of the mutations found in BTK result in a total absent BTK protein expression, leading to a block in B cell differentiation. However, some BTK mutations do not affect BTK expression completely and are generally associated with a milder phenotype. (31, 32) Next to these variable phenotypes, other mutations within the signaling cascade of BTK phosphorylation have been shown to cause defects. Agammaglobulinemia and absence of B cells have been described in a patient lacking the p85α subunit of PI3K. (33)

Since defects in early B cell development lead to the absence of B cells or their severely decreased number in the blood, functional B cell studies cannot be performed and hence do not represent an important part in this thesis.

Figure 1. Showing seven distinct groups, with genetic defects associated, that can be recognized in defective

B cell development and differentiation. Namely defects in early B cell development, early survival, activation, T cell dependent stimulation (TD), T cell independent stimulation (TI), class switch recombination (CSR) and late survival. (Adapted from (58))

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1 CLASS-SWITCH RECOMBINATION DEFECTS (CSR)

A very rare group of PADs are the so-called hyper-IgM syndromes (HIGM) or class-switch recombination (CSR) defects, which are characterized by a low-to-normal or elevated serum IgM in the absence of any other immunoglobulin (Ig) isotypes in the serum. A number of unique gene defects (e.g. in CD40L, CD40, AICDA, UNG, and IKBKG) have been identified that lead to a defect in the CSR mechanism required for the transition of the initial low-affinity IgM to an Ig isotype of increased affinity.

COMMON VARIABLE IMMUNODEFICIENCY (CVID)

The most prevalent of the PAD diagnoses is common variable immunodeficiency (CVID). The etiology of CVID seems to have a complex genetic cause, although few monogenetic mutations may explain a CVID-like phenotype (in e.g. TACI, ICOS, CD19, CD20 and BAFFR). CVID is diagnosed by serum IgG levels below 2 SD of the normal range in the presence of decreased IgA and/or IgM levels, recurrent infections, impaired immunization responses, an age above two years and exclusion of other causes of hypogammaglobulinemia. These criteria were set by the “European Society for Immunodeficiencies (ESID)”. (28, 34) Early detection is shown to be of great importance, since starting treatment earlier in life gives superior survival. (27) Most cases are diagnosed in young adulthood, although symptoms can start much earlier in life. (35) Some of these patients are first diagnosed with a hypogammaglobulinemia or (combined) IgG-subclass deficiency but do not fulfill all the criteria for CVID and are therefore called “CVID-like”, which may or may not evolve over time to become full-blown CVID. (36)

Over the years many groups have worked on classifying CVID based on different parameters, e.g. B cell phenotype, clinical presentation and B cell function. (37-40) The most comprehensive and broadly used classification to date is the CVID-EUROclass 2008, discriminating between patients with nearly absent B cells (less than 1%), severely reduced switched memory B cells (less than 2%),

and expansion of transitional (more than 9%) or CD21low_{B cells (more than 10%) of the circulating B}

cells(41) (Figure 2). Classification may lead to better outcome, earlier detection and possibly finding underlying causative defects.

OTHERS SPECIFIC COMBINED IMMUNE DEFECTS INVOLVING

B CELLS DYSFUNCTION

An example of defects in migration is shown by deficiency in a cytoskeleton regulator, dedicator of cytokinesis 8 (DOCK8). Patients are described to have a defect in the generation of marginal zone B cells and impairment in the retention of B cells at the germi nal centre as well as BCR affinity maturation. Resulting in low IgM levels, variable IgG antibody responses, low num bers of memory B cells and defective B cell responses to Toll-like receptor (TLR) activation. After ligation of TLR9, DOCK8 linked TLR9 to a Src–Syk–STAT3 cascade essential for B cell proliferation and differentiation, thus making DOCK8 function as an adaptor in a TLR9-MyD88 signaling pathway in B cells. It remains unclear if these defects are solely the result of B cell defects since T cells also a defective function. (42-44)

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Crucial for B cell activation is the canonical nuclear factor κB (NFκB) pathway, in which

numerous defects are described. B cell activation following BCR and CD40 triggers this NFκB pathway, via activation of IκB kinase (IKK) complex, consisting of IKKα, IKKβ and NEMO (also known

as IKKγ). Defects are described in IKBKB (encoding IKKβ) and IKBKG (encoding NEMO), in which

hypomorphic mutations in the gene for NEMO in most of the cases will lead to immunodeficiency together with anhydrotic ectodermal dysplasia (EDA) (45, 46).

Another example of combined immune defects involving B cells dysfunction is a defect in the scaffolding protein CARD11, essential for linking the BCR with the T cell receptor (TCR) and also

activating the NFκB pathway. Defects in CARD11 lead to agammaglobulinaemia in the presence

of a normal number of B cells, although the B cell fraction in the blood can show an increased number of transitional B cells. (47, 48) Similar defects in the NFκB pathway have been described for the Bcl-10- and MALT-1-deficient individuals that have been reported thus far, who present with defective T and B cell activation following stimulation via their antigen receptors. As a result a completely naïve B cell phenotype is found without specific antibody responses and most often a profound hypogammaglobulinemia. (49)

Recently heterozygous NFKB1 and NFKB2 defects have been reported. Both NFKB1 (50) and

NFKB2 (51, 52) can present as a predominantly humoral immunodeficiency with or without signs

Figure 2. Showing CVID-EUROclass 2008, discriminating between patients with nearly absent B cells

(less than 1%), severely reduced switched memory B cells (less than 2%), and expansion of transitional (more than 9%) or CD21low_{B cells (more than 10%) of the circulating B cells.}

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of autoimmunity, essentially similar to CVID while the causative signaling molecules may be

ubiquitously expressed – as is true for the NFκB p50 protein as encoded by NFKB1.

As described above a deficiency was identified in p85alpha subunit of PI3K that results in agammaglobulinemia by an early block in B cell development caused by a PIK3R1 gene defect. (53) More recently, constitutively active PI3K signaling was described as the Activated PI3Kδ Syndrome (APDS) (54-57) caused by autosomal-dominant gene defects, which result in a combined B and T cell dysfunction.

AIM AND OUTLINE OF THIS THESIS

In most patients with PAD the underlying pathophysiological mechanism is unknown and no monogenetic cause can be found. Treatment consisting of replacement immunoglobulins with or without antibiotic prophylaxis (or even immunosuppressive medication to reduce granuloma formation if present), can be started depending on clinical history, symptoms and co-existing organ damage. Antibody replacement therapy and additional protective measures should be decided upon, independently of the known or unidentified causative genetic defects, to improve long term outcomes.

Gaining knowledge of the pathophysiology and underlying genetic defects is still of great value. Next to the intrinsic need to know, we can use these patients to better understand normal B cell development and differentiation, designing diagnostic tests for earlier diagnosis and possibly better treatment option.

We introduce a B cell differentiation assay, in which we only use a small amount of peripheral blood mononuclear cells (PBMC). This assay was tested for use in a diagnostic setting, and reference values for all ages have been generated. This provided a way to study B cell differentiation of cells from healthy donors or patients with known or unknown B cell defects.

The aims of the studies described in this thesis were to improve the knowledge of B cell development and differentiation, and to improve detection, diagnosis and classification of PADs in children and adults.

In chapter 2 we described our assay providing an in-vitro way to study B cell differentiation. By mimicking either T cell-dependent or T cell-independent activation we showed clear formation of immunoglobulin-producing PBs. Along with introducing this new method, age-defined reference ranges were created, showing big differences in B cell function between age groups.

In chapter 3 we used our assay to describe a subgroup of CVID patient who have a functional CSR defect. We compared them to patients with known, genetically proven, CSR defects. This added a new subgroup of patients, with specific B cell defects, within the commonly used CVID classifications.

In chapter 4 we adapted our B cell differentiation assay to be used in a high throughput compound screen to test for the effect of various small molecules on B cell survival, proliferation and differentiation. We aimed to gain insight in previously unknown pathways important for B cell development, potentially leading to new treatment approaches for PAD or autoimmunity. The data showed previously unknown B cell specific effects of mTOR inhibitors on human B cell activation and differentiation.

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provided data showing that we were not able to reproduce previously published data, questioning In chapter 5 we describe the controversy around the existence of B helper neutrophils. We

the used methods and therefore validity of these data.

In chapter 6 & 7 our B cell differentiation assay was used to report on two patients with PID in highly informative case reports. Chapter 6 describes a patient with a DOCK8 deficiency, causing autosomal-recessive Hyper-IgE syndrome (HIES), showing almost absent B cell function. Chapter

7 describes a novel hypomorphic mutation in the gene IL2RG, encoding the common γ-chain

CD132 for several interleukin receptors, causing in the index case a form of X-CID (i.e. a form of very late-onset SCID). B cell function is clearly defective, showing the necessity of a functional

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32. Bykowsky, M. J., R. N. Haire, Y. Ohta, H. Tang, S. S. Sung, E. S. Veksler, J. M. Greene, S. M. Fu, G. W. Litman, and K. E. Sullivan. 1996. Discordant phenotype in siblings with X-linked agammaglobulinemia. Am J Hum Genet 58: 477-483.

33. Al-Herz, W., A. Bousfiha, J. L. Casanova, T. Chatila, M. E. Conley, C. Cunningham-Rundles, A. Etzioni, J. L. Franco, H. B. Gaspar, S. M. Holland, C. Klein, S. Nonoyama, H. D. Ochs, E. Oksenhendler, C. Picard, J. M. Puck, K. Sullivan, and M. L. Tang. 2014. Primary immunodeficiency diseases: an update on the classification from the international union of immunological societies expert committee for primary immunodeficiency. Frontiers in immunology 5: 162.

34. International Union of Immunological Societies Expert Committee on Primary, I., L. D. Notarangelo, A. Fischer, R. S. Geha, J. L. Casanova, H. Chapel, M. E. Conley, C. Cunningham-Rundles, A. Etzioni, L. Hammartrom, S. Nonoyama, H. D. Ochs, J. Puck, C. Roifman, R. Seger, and J. Wedgwood. 2009. Primary immunodeficiencies: 2009 update. The Journal of allergy and clinical immunology 124: 1161-1178.

35. Quinti, I., A. Soresina, G. Spadaro, S. Martino, S. Donnanno, C. Agostini, P. Claudio, D. Franco, A. Maria Pesce, F. Borghese, A. Guerra, R. Rondelli, A. Plebani, and N. Italian Primary Immunodeficiency. 2007. Long-term follow-up and outcome of a large cohort of patients with common variable immunodeficiency. Journal of clinical immunology 27: 308-316.

36. Brignier, A. C., N. Mahlaoui, C. Reimann, C. Picard, S. Kracker, N. de Vergnes, F. Rieux-Laucat, P. Frange, F. Suarez, B. Neven, A. Masseau, N. Aladjidi, J. Donadieu, A. Corby, B. Bienvenu, P. Cony-Makhoul, A. Fischer, M. Cavazzana, and A. Durandy. 2015. Early-onset hypogammaglobulinemia: A survey of 44 patients. The Journal of allergy and clinical immunology 136: 1097-1099 e1092.

37. Rosel, A. L., C. Scheibenbogen, U. Schliesser, A. Sollwedel, B. Hoffmeister, L. Hanitsch, H. von Bernuth, R. Kruger, K. Warnatz, H. D. Volk, and S. Thomas. 2014. Classification of common variable immunodeficiencies using flow cytometry and a memory B-cell functionality assay. The Journal of allergy and clinical immunology.

38. Schatorje, E. J., E. F. Gemen, G. J. Driessen, J. Leuvenink, R. W. van Hout, M. van der Burg, and E. de Vries. 2011. Age-matched reference values for B-lymphocyte subpopulations and CVID classifications in children. Scandinavian journal of immunology 74: 502-510.

39. van de Ven, A. A., L. van de Corput, C. M. van Tilburg, K. Tesselaar, R. van Gent, E. A. Sanders, M. Boes, A. C. Bloem, and J. M. van Montfrans. 2010. Lymphocyte characteristics in children with common variable immunodeficiency. Clinical immunology 135: 63-71.

40. Warnatz, K., and M. Schlesier. 2008. Flowcytometric phenotyping of common variable immunodeficiency. Cytometry. Part B, Clinical cytometry 74: 261-271.

41. Wehr, C., T. Kivioja, C. Schmitt, B. Ferry, T. Witte, E. Eren, M. Vlkova, M. Hernandez, D. Detkova, P. R. Bos, G. Poerksen, H. von Bernuth, U. Baumann, S. Goldacker, S. Gutenberger, M. Schlesier, F. Bergeron-van der Cruyssen, M. Le Garff, P. Debre, R. Jacobs, J. Jones, E. Bateman, J. Litzman, P. M. van Hagen, A. Plebani, R. E. Schmidt, V. Thon, I. Quinti, T. Espanol, A. D. Webster, H. Chapel, M. Vihinen, E. Oksenhendler, H. H. Peter, and K. Warnatz. 2008. The EUROclass trial: defining subgroups in common variable immunodeficiency. Blood 111: 77-85.

42. Zhang, Q., J. C. Davis, I. T. Lamborn, A. F. Freeman, H. Jing, A. J. Favreau, H. F. Matthews, J. Davis, M. L. Turner, G. Uzel, S. M. Holland, and H. C. Su. 2009. Combined immunodeficiency associated with DOCK8 mutations. The New England journal of medicine 361: 2046-2055.

43. Randall, K. L., T. Lambe, A. L. Johnson, B. Treanor, E. Kucharska, H. Domaschenz, B. Whittle, L. E. Tze, A. Enders, T. L. Crockford, T. Bouriez-Jones, D. Alston, J. G. Cyster, M. J. Lenardo, F. Mackay, E. K. Deenick, S. G. Tangye, T. D. Chan, T. Camidge, R. Brink, C. G. Vinuesa, F. D. Batista, R. J. Cornall, and C. C. Goodnow. 2009. Dock8 mutations cripple B cell immunological synapses, germinal centers and long-lived antibody production. Nature immunology 10: 1283-1291.

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44. Jabara, H. H., D. R. McDonald, E. Janssen, M. J. Massaad, N. Ramesh, A. Borzutzky, I. Rauter, H. Benson, L. Schneider, S. Baxi, M. Recher, L. D. Notarangelo, R. Wakim, G. Dbaibo, M. Dasouki, W. Al-Herz, I. Barlan, S. Baris, N. Kutukculer, H. D. Ochs, A. Plebani, M. Kanariou, G. Lefranc, I. Reisli, K. A. Fitzgerald, D. Golenbock, J. Manis, S. Keles, R. Ceja, T. A. Chatila, and R. S. Geha. 2012. DOCK8 functions as an adaptor that links TLR-MyD88 signaling to B cell activation. Nature immunology 13: 612-620.

45. Zonana, J., M. E. Elder, L. C. Schneider, S. J. Orlow, C. Moss, M. Golabi, S. K. Shapira, P. A. Farndon, D. W. Wara, S. A. Emmal, and B. M. Ferguson. 2000. A novel X-linked disorder of immune deficiency and hypohidrotic ectodermal dysplasia is allelic to incontinentia pigmenti and due to mutations in IKK-gamma (NEMO). Am J Hum Genet 67: 1555-1562.

46. Hanson, E. P., L. Monaco-Shawver, L. A. Solt, L. A. Madge, P. P. Banerjee, M. J. May, and J. S. Orange. 2008. Hypomorphic nuclear factor-kappaB essential modulator mutation database and reconstitution system identifies phenotypic and immunologic diversity. The Journal of allergy and clinical immunology 122: 1169-1177 e1116.

47. Stepensky, P., B. Keller, M. Buchta, A. K. Kienzler, O. Elpeleg, R. Somech, S. Cohen, I. Shachar, L. A. Miosge, M. Schlesier, I. Fuchs, A. Enders, H. Eibel, B. Grimbacher, and K. Warnatz. 2013. Deficiency of caspase recruitment domain family, member 11 (CARD11), causes profound combined immunodeficiency in human subjects. The Journal of allergy and clinical immunology 131: 477-485 e471.

48. Snow, A. L., W. Xiao, J. R. Stinson, W. Lu, B. Chaigne-Delalande, L. Zheng, S. Pittaluga, H. F. Matthews, R. Schmitz, S. Jhavar, S. Kuchen, L. Kardava, W. Wang, I. T. Lamborn, H. Jing, M. Raffeld, S. Moir, T. A. Fleisher, L. M. Staudt, H. C. Su, and M. J. Lenardo. 2012. Congenital B cell lymphocytosis explained by novel germline CARD11 mutations. The Journal of experimental medicine 209: 2247-2261.

49. Perez de Diego, R., S. Sanchez-Ramon, E. Lopez-Collazo, R. Martinez-Barricarte, C. Cubillos-Zapata, A. Ferreira Cerdan, J. L. Casanova, and A. Puel. 2015. Genetic errors of the human caspase recruitment domain-B-cell lymphoma 10-mucosa-associated lymphoid tissue lymphoma-translocation gene 1 (CBM) complex: Molecular, immunologic, and clinical heterogeneity. The Journal of allergy and clinical immunology 136: 1139-1149. 50. Fliegauf, M., V. L. Bryant, N. Frede, C. Slade, S. T. Woon, K. Lehnert, S. Winzer, A. Bulashevska, T. Scerri, E.

Leung, A. Jordan, B. Keller, E. de Vries, H. Cao, F. Yang, A. A. Schaffer, K. Warnatz, P. Browett, J. Douglass, R. V. Ameratunga, J. W. van der Meer, and B. Grimbacher. 2015. Haploinsufficiency of the NF-kappaB1 Subunit p50 in Common Variable Immunodeficiency. Am J Hum Genet 97: 389-403.

51. Chen, K., E. M. Coonrod, A. Kumanovics, Z. F. Franks, J. D. Durtschi, R. L. Margraf, W. Wu, N. M. Heikal, N. H. Augustine, P. G. Ridge, H. R. Hill, L. B. Jorde, A. S. Weyrich, G. A. Zimmerman, A. V. Gundlapalli, J. F. Bohnsack, and K. V. Voelkerding. 2013. Germline mutations in NFKB2 implicate the noncanonical NF-kappaB pathway in the pathogenesis of common variable immunodeficiency. Am J Hum Genet 93: 812-824.

52. Brue, T., M. H. Quentien, K. Khetchoumian, M. Bensa, J. M. Capo-Chichi, B. Delemer, A. Balsalobre, C. Nassif, D. T. Papadimitriou, A. Pagnier, C. Hasselmann, L. Patry, J. Schwartzentruber, P. F. Souchon, S. Takayasu, A. Enjalbert, G. Van Vliet, J. Majewski, J. Drouin, and M. E. Samuels. 2014. Mutations in NFKB2 and potential genetic heterogeneity in patients with DAVID syndrome, having variable endocrine and immune deficiencies. BMC medical genetics 15: 139.

53. Conley, M. E., A. K. Dobbs, A. M. Quintana, A. Bosompem, Y. D. Wang, E. Coustan-Smith, A. M. Smith, E. E. Perez, and P. J. Murray. 2012. Agammaglobulinemia and absent B lineage cells in a patient lacking the p85alpha subunit of PI3K. The Journal of experimental medicine 209: 463-470.

54. Deau, M. C., L. Heurtier, P. Frange, F. Suarez, C. Bole-Feysot, P. Nitschke, M. Cavazzana, C. Picard, A. Durandy, A. Fischer, and S. Kracker. 2015. A human immunodeficiency caused by mutations in the PIK3R1 gene. The Journal of clinical investigation 125: 1764-1765.

55. Angulo, I., O. Vadas, F. Garcon, E. Banham-Hall, V. Plagnol, T. R. Leahy, H. Baxendale, T. Coulter, J. Curtis, C. Wu, K. Blake-Palmer, O. Perisic, D. Smyth, M. Maes, C. Fiddler, J. Juss, D. Cilliers, G. Markelj, A. Chandra, G. Farmer, A. Kielkowska, J. Clark, S. Kracker, M. Debre, C. Picard, I. Pellier, N. Jabado, J. A. Morris, G.

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Barcenas-Morales, A. Fischer, L. Stephens, P. Hawkins, J. C. Barrett, M. Abinun, M. Clatworthy, A. Durandy, R. Doffinger, E. R. Chilvers, A. J. Cant, D. Kumararatne, K. Okkenhaug, R. L. Williams, A. Condliffe, and S. Nejentsev. 2013. Phosphoinositide 3-kinase delta gene mutation predisposes to respiratory infection and airway damage. Science 342: 866-871.

56. Crank, M. C., J. K. Grossman, S. Moir, S. Pittaluga, C. M. Buckner, L. Kardava, A. Agharahimi, H. Meuwissen, J. Stoddard, J. Niemela, H. Kuehn, and S. D. Rosenzweig. 2014. Mutations in PIK3CD can cause hyper IgM syndrome (HIGM) associated with increased cancer susceptibility. Journal of clinical immunology 34: 272-276. 57. Lucas, C. L., H. S. Kuehn, F. Zhao, J. E. Niemela, E. K. Deenick, U. Palendira, D. T. Avery, L. Moens, J. L.

Cannons, M. Biancalana, J. Stoddard, W. Ouyang, D. M. Frucht, V. K. Rao, T. P. Atkinson, A. Agharahimi, A. A. Hussey, L. R. Folio, K. N. Olivier, T. A. Fleisher, S. Pittaluga, S. M. Holland, J. I. Cohen, J. B. Oliveira, S. G. Tangye, P. L. Schwartzberg, M. J. Lenardo, and G. Uzel. 2014. Dominant-activating germline mutations in the gene encoding the PI(3)K catalytic subunit p110delta result in T cell senescence and human immunodeficiency. Nature immunology 15: 88-97.

58. Berkowska, M. A., M. van der Burg, J. J. van Dongen, and M. C. van Zelm. 2011. Checkpoints of B cell differentiation: visualizing Ig-centric processes. Annals of the New York Academy of Sciences 1246: 11-25.

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Identification of B-cell defects

using age-defined reference ranges

for in-vivo and in-vitro

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2 ABSTRACT

Background

Primary immunodeficiencies consist to a large extent of B-cell defects, as indicated by inadequate antibody levels or response upon immunization. Many B-cell defects have not yet been well characterized.

Objective

To create reliable in-vivo and in-vitro assays to routinely analyze human B-cell differentiation, proliferation, and immunoglobulin production and to define reference ranges for different age categories. The in-vitro assays were applied to classify the developmental and/or functional B-cell defects in patients previously diagnosed with common variable immunodeficiency (CVID).

Methods

Apart from standard immunophenotyping of circulating human B-cell subsets, an in-vitro CFSE dilution assay was used for the assessment of proliferative capacity comparing T-cell-dependent and T-cell-independent B-cell activation. Plasmablast/plasmacell differentiation was assessed by staining for CD20, CD38 and CD138, and measurement of in-vitro immunoglobulin secretion.

Results

At young age, B-cells proliferate upon in-vitro activation but neither differentiate nor produce IgG. These latter functions reached ‘adult’ levels at 5 and 10 years of age for T-cell-dependent versus T-cell-independent stimulations, respectively. The capacity of B-cells to differentiate into plasmablasts and to produce IgG appeared to be contained within the switched memory B-cell pool. Using these assays, we could categorize CVID patients into subgroups and identified a class-switch recombination defect caused by an UNG mutation in one of the patients.

Conclusion

We defined age-related reference ranges for human B-cell differentiation. Our findings indicate that

in-vivo B-cell functionality can be tested in-vitro and help to diagnose suspected B-cell defects.

Daan J. aan de Kerk, MD 1, 2_{, Machiel H. Jansen, MSc}1, 2_{, Ineke J.M. ten Berge, MD PhD}3_,

Ester M.M. van Leeuwen, PhD 2_{, Taco W. Kuijpers, MD PhD}1

Journal of Immunology, 2013 May; 190(10): 5012-9

1._{Emma Children’s Hospital, Academic Medical Center (AMC), Amsterdam, The Netherlands} 2._{Department of Experimental Immunology, AMC, Amsterdam, The Netherlands}

3._{Renal Transplant Unit, Department of Nephrology, Division of Internal Medicine, Academic Medical Centre,}

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2 INTRODUCTION

In the Western world about 1 in 500 adults have a primary immunodeficiency (PID)(1-3). About 65% of PID cases show altered antibody production which can be part of a combined T- and B-cell defect or an B-cell-intrinsic defect. Common variable immunodeficiency (CVID) is the most prevalent serious PID diagnosis with an antibody deficiency caused by B-cell dysfunction. Although a few genetic mutations may contribute (e.g. TACI, ICOS, CD19, CD20 and BAFFR), the etiology is unknown in most cases(3). Twenty-fold less common is the Hyper-IgM syndrome (HIGM), characterized by a low, normal or elevated serum IgM in the absence of other immunoglobulin (Ig) isotypes, as defined by unique genetic defects (e.g. CD40L, CD40, AICDA, UNG, IKKG) that all lead to a defect in Ig class-switch recombination (CSR)(4,5). Finally, agammaglobulinemia patients usually lack circulating B-cells, of which most males suffer from X-linked Bruton’s disease caused by mutations in BTK(6).

Many B-cell defects still have not been characterized in functional terms and at the genetic level. Hence, we need good tools to classify at which developmental stage such B-cell defect can be located. For this, we have developed a cell culture system to determine the capacity of mature peripheral blood B-cells to proliferate and differentiate into activated B-cells and Ig-secreting plasmablasts (PBs) and plasmacells (PCs). Upon triggering by antigen, B-cells normally become activated, expand and differentiate from naïve B-cells into memory B-cells or antibody-secreting cells. Differentiation may proceed largely via two distinct pathways, depending on the nature of the stimulus, i.e. T-cell-dependent antigens (typically protein structures) or a T-cell-independent antigen (typically carbohydrates).

T-cell-dependent differentiation leads to long-lived, highly specific, immunoglobulin-producing PCs. Activation results in the formation of germinal centers in lymphoid organs such as the lymph

nodes, spleen and Peyer’s patches(7), where activated CD4+_{T-cells interact with activated B-cells}

through direct CD40L-CD40 interactions, and the release of soluble factors such as IL-21. These T-B-cell interactions are essential for somatic hypermutation (SHM) and CSR in B-cells, both needed to induce differentiation into high-affinity IgG-producing PCs(7,8). T-cell-independent activation most often leads to short-lived, moderate-affinity Ig-producing cells. When activated in a T-cell-independent manner, activated B-cells mostly differentiate into IgM-producing PBs(9,10), as demonstrated for the polyclonal activation either by B-cell-receptor (BcR)-independent Toll-Like Receptor (TLR) activation or BcR-dependent carbohydrate antigens(11,12).

When screening for immune defects in children, the healthy controls used mainly consist of adults. However, the composition of the human B-cell compartment changes greatly during childhood and there is considerable variability in the size of the adult memory B-cell pool due to antigen exposure during life which will influence the test results(13).

Using an in-vitro culture system to investigate B-cell differentiation in humans, we generated reference ranges which demonstrated clear age-dependent differences in the acquisition of immunophenotypic and functional capacity to differentiate into PBs and PCs. Screening a cohort of 15 patients diagnosed with or suspected of CVID, we determined the presence of subgroups among these CVID patients, while narrowing the search for any underlying monogenic defect. In further characterization of these CVID patients we identified a rare UNG mutation causing his B-cell defect. In addition to other molecular techniques, using replication history or extensive immunophenotyping(14,15), our functional assay forms a helpful tool in the evaluation of B-cell

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2

differentiation stages and contributes to a more precise characterization of patients suspected of humoral PID.

MATERIALS & METHODS

Samples

Heparinized peripheral blood samples were collected from healthy control individuals as well as patients for routine diagnostics. Blood samples from healthy age-matched control donors were collected as rest material of blood drawn for reasons unrelated to the study, as approved by the local medical ethical committee.

PBMCs were isolated using standard density gradient centrifugation techniques using Lymphoprep (Nycomed, Oslo, Norway) and stored in liquid nitrogen until use. In some of the experiments, the patient’s samples were analyzed simultaneously with PBMCs from one or two age-matched controls.

Flowcytometry

PBMCs were resuspended in PBS, containing 0.5%(w/v) BSA and 0.01% sodium azide. PBMCs were incubated with saturating concentrations of fluorescently labeled conjugated MoAbs. For intracellular staining, surface staining was performed first, cells were then fixed and permeabilized with Cytofix/Cytoperm and Perm/Wash solutions (BD-Biosciences, Erembodegem, Belgium) and subsequently stained for intracellular markers. Analysis of cells was performed using either a FACSCalibur flowcytometer and CellQuest software or a FACSCanto-II flowcytometer and FACSDiva software (BD-Biosciences). Absolute numbers of T-cells and B-cells were determined with Multitest six-color reagents (BD Biosciences), according to the manufacturer’s instructions. The following directly conjugated monoclonal antibodies (MoAbs) were used for flowcytometry: CD4-PE, CD8-PerCP-Cy-5.5, CD3-APC, CD19-PerCP-Cy 5.5, CD19 Alexa-700, APC, CD20-PerCP-Cy 5.5, CD38 PE-Cy7, CD132-biotin, CD138 APC and IgG-PE from BD-biosciences, CD27-FITC from Sanquin (Amsterdam, The Netherlands), IgM-PE (ITK-diagnostics, Uithoorn, the Netherlands).

B-cell activation in-vitro

PBMCs were resuspended in PBS at a concentration of 5–10×106_{cells/ml and labelled with 0.5μM}

CFSE (Molecular Probes) in PBS for 10 minutes at 37°C under constant agitation. Cells were washed and subsequently resuspended in IMDM supplemented with 10% fetal calf serum (BioWhittaker), antibiotics, and 3.57×10–4%(v/v) β-mercapto-ethanol (Merck). Labelled PBMCs containing a fixed

number of B cells (×105_{per well) were cultured in 48-well flat-bottomed plates for 6 days at 37°C and}

stimulated with saturating amounts of anti-IgM mAb (clone MH15; Sanquin), anti-CD40 mAb (clone 14G7; Sanquin), 2ng/ml IL-21 (Invitrogen), or 200μg/ml CpG oligodeoxynucleotide 2006 (Invivogen), with 50U/ml IL-2 (R&D Systems).

Proliferation of the B cells was assessed by measuring CFSE dilution. The precursor frequency (percentage of cells in the initial population that underwent one or more divisions after culture)

was calculated as follows: [Σn≥1(Pn/2n)]/[Σn≥0(Pn/2n)], where n is the division number that cells

have gone through and Pn is the number of cells in division n. The mean number of divisions of

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2 IgG & IgM ELISA

Supernatants were tested for secreted IgM and IgG with an in-house ELISA using polyclonal rabbit anti-human IgG and IgM reagents and a serum protein calibrator all from Dako (Heverlee, Belgium)(16).

Statistics

Differences between immunoglobulin levels were calculated by 2-sided, 2-tailed Student’s t-test. For correlations, the Spearman nonparametric correlation test was used. P less than .05 was considered statistically significant.

RESULTS

Age-dependent distribution of B-cell differentiation during childhood

During childhood the composition of the human B-cell compartment changes from a very naive

to a more differentiated phenotype, coinciding with the appearance of a CD27+_{memory B-cell}

compartment. Using surface IgD and CD27 expression, different differentiation stages of mature

peripheral B-cells can be distinguished, discriminating naïve (IgD+_/CD27−_{), non-switched (IgD}+_/

CD27+_{) and switched memory (IgD}−_/CD27+_{) B-cells(17,18).}

In the adult PBMC fraction, the B-cell phenotype demonstrated the presence of a clear memory B-cell compartment which is missing in cord-blood PBMCs where all B-cells are naïve (Figure 1A). A large number of healthy pediatric controls aged 0-18 years of age (n=315 in total) were tested over the last 2 years and divided into distinct age subcategories, in addition to adult controls. The amount of naïve B-cells decreased with age during childhood (Figure 1B), whereas non-switched and switched memory B-cells increased accordingly. There is considerable variation within each age groups which may be explained by variation in the genetic background, racial differences, antigen exposure and many other unknown factors that may contribute to B-cell maturation.

The effect of B-cell activation on differentiation in culture

Using the same age categories, we determined the capacity of B-cells to differentiate in-vitro. This was tested in various ways. First, T-cell-dependent B-cell stimulation was mimicked by the combinations of αIgM/αCD40 or αIgM/αCD40/IL-21, in which the surface-IgM receptor as

B-cell receptor (BcR) on the majority of naïve B-cells was activated together with αCD40 and IL-21

to simulate T-cell help(19).

Secondly, T-cell-independent B-cell activation was tested with CpG in the absence or presence of IL-2. In this case, CpG stimulates B-cells through TLR9 without BcR triggering while IL-2 adds to the effect by direct B-cell activation(20).

After 6 days of culture the phenotype of B-cells from healthy adults were analyzed (Figure 2A).

Without cell activation no increase in memory B-cell numbers (IgD−_CD27+_{) was observed. Stimulation}

with αIgM/αCD40 gave rise to a consistent but minor fraction of memory B-cells, whereas

the addition of IL-21 resulted in markedly increased differentiation into a discrete population of

highly activated memory B-cells (IgD−_CD27high_{). Also the T-cell-independent activation of B-cells}

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2

Figure 1. Age-dependent changes of B-cell differentiation during childhood. (A) B-cell subsets of representative

blood samples from a healthy adult control and a healthy cord-blood, numbers indicate percentages in the corresponding quadrant. Gated on CD19+_CD20+_{lymphocytes. (B) Quantification of B-cell subsets of healthy}

pediatric controls (n=315) and adults.

of culture. This fraction of CD27high_{memory B-cells has been previously shown to be committed to}

the plasmacell lineage(21).

Under the same culture conditions, B-cell proliferation was tested by the analysis of CFSE dilution. Upon B-cell activation, the appearance of two specific subsets of Ig-producing B-cells

became evident (Figure 2B): i.e. plasmablasts (PBs; CD38bright_/CD20dull_{), and plasmacells (PCs; CD138}+_/

CD38bright_/CD20dull)(7). T-cell-dependent B-cell activation by αIgM/αCD40 induced proliferation but

failed to induce differentiation into PBs or PCs. Of various cytokines (i.e. IL-2, IL-4, IL-15 or IL-21) tested, IL-21 was most potent in inducing additional B-cell proliferation (data not shown). CpG induced strong B-cell proliferation and PB/PC formation to effect cytokines such as IL-2 added only slightly (Figure 2B).

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2

Subtyping by flowcytometry indicated that the PB/PC phenotype was largely contained within

the CD27high_{and CD20}dull _{B-cell population (Supplementary Figure 1)(21). Noticeably, almost half of}

the CD27high_{B-cell population in culture had not undergone any cell division at all, while being to}

a large extent also CD20dull_{and CD138}+_{, suggesting that some B-cells can differentiate into PBs/PCs}

without proliferation.

Age-dependent changes in B-cell differentiation

Age dependent capacity of B-cells to proliferate and differentiate into PBs/PCs was tested in healthy pediatric controls of different age categories. As shown in Figure 3A, differentiation into PBs steadily increased with age. The lack of B-cell differentiation at young age was not the result of a difference in the ability to proliferate. In fact, cord-blood B-cells had the highest precursor frequency (Supplementary Figure 2A).

When summarizing multiple experiments (Figure 3B/3C; n=3-5), the in-vitro differentiation into

PBs and PCs (CD38bright _{and CD138}+_{, respectively) was found to increase over time during childhood,}

which was most apparent in the CFSElow_{B-cell fraction. Unstimulated cells did not show any CD38}bright

cells. This was also true for αIgM/αCD40-activated B-cells, which need addition of IL-21 to induce

CD38bright_{PBs depending on age. Stimulation with CpG alone gave rise to CD38}bright _{cells from the age}

of about 5 years onward. Addition of IL-2 resulted in the appearance of CD38bright _{cells at younger}

age, coinciding with the ability to differentiate into CD138+_PCs.

Although age-dependent differences between T-cell-independent and T-cell-dependent responses may relate to the expression levels of their main (co)activating receptors, cord-blood B-cells were found to have only slightly lower levels of intracellular TLR9 expression compared to adult B-cells (Supplementary Figure 2B). When analyzing the IL21R expression, no differences between cord-blood and adult naïve B-cells were observed, whereas IL21R expression on adult memory B-cells from the same donor was lower than on the naïve B-cells (Supplementary Figure 2C).

Age-dependent changes in production and release of immunoglobulins

To investigate whether the in-vitro induced PBs/PCs were able to release immunoglobulins, we measured IgG and IgM in the supernatants of the 6-day cultures (Figures 4A/B).

Upon αIgM/αCD40 stimulation the B-cells produced no IgG. Addition of IL-21 to these αIgM/ αCD40 conditions resulted in the production of IgG, starting at infancy and steadily increasing to levels comparable to adult levels from 5-10 yrs of age onward. Upon stimulation with CpG alone, B-cells from controls in the youngest age categories did not release IgG. From the age of 5 years onward the supernatant contained increasing levels of IgG (in contrast to T-cell-dependent stimulation by αIgM/αCD40). Adding IL-2 increased IgG production, in particular at young age. However, at this young age PBs and PCs still constitute a small fraction of total B-cell number (Figure 3A).

IgM production was simultaneously determined (Figure 4B), except for conditions where αIgM was used for B-cell stimulation. T-cell-independent stimulation with CpG gave a similar pattern as observed for IgG over time, with the remarkable exception of IgM production in

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Figure 2. Differentiation of human B-cells upon activation. The capacity of B-cells, from a healthy adult control,

to proliferate and differentiate in-vitro was tested. CFSE labeled PBMCs were cultured for 6 days, normalized for B-cell numbers (1*105_{B-cells/ well). T-cell-dependent B-cell stimulation was mimicked by the combinations of}

αIgM/αCD40 or αIgM/αCD40/IL-21. T-cell-independent B-cell activation was tested with CpG in the absence or presence of IL-2. (A) Representative FACS plots of B-cell subset distribution after 6 days of culture in the presence of the indicated stimuli. Gated on CD19+_{lymphocytes. (B) Representative FACS plots showing}

CFSE dilution, indicating proliferation, after 6 days of culture and the emergence of two specific subsets of Ig-producing B-cells Plasmablasts (CD38bright_{) and Plasmacells (CD138}+_).

blood samples. This IgM production at birth had completely disappeared in the infant samples (0.5-1 year of age), although addition of IL-2 to CpG helped to restore IgM production at very young age. The data collectively show that, apart from cord-blood samples, the production of IgG and IgM correlates with the formation of PBs/PCs in-vitro and thus depends on age.

Activation and reactivity depend on B-cell differentiation

Whether the inability of B-cells to differentiate in-vitro into PBs/PCs at young age is due to cell-intrinsic mechanisms or depends on the lack of circulating memory B-cells in-vivo, was addressed. First, the amount of IgG produced after culture showed a positive correlation with the number

of circulating memory B-cells (IgD−_CD27+_{) at the start of our cultures (Supplementary Figure 3).}

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Figure 3. Age-dependent changes in B-cell differentiation. Proliferative capacity and simultaneous induction of

the differentiation program in human B-cells was tested in healthy pediatric controls of different age categories and adults. Culturing lymphocytes normalized for B-cell numbers for 6 days upon activation as described in the Methods section. (A) Representative examples from healthy children from different age categories are shown. Proliferation measured by CFSE dilution and differentiation into CD38bright_{plasmablasts are shown.}

Gated on CD19+_CD20+_{lymphocytes. (B/C) Percentage of PB and PC are shown for multiple experiments (N=3-5).}

Discriminating between divided (CFSElow_{) and undivided (CFSE}high_{) cells, CD38}bright _PBs_{are shown in (B) and}

CD138+_{PCs in (C).}

with or without the addition of autologous T-cells, i.e. non-B-cells (CD19−_CD20−_{) isolated from}

the same blood sample, and activated as in our previous experiments (Figure 5A).

After sorting, stimulation of naïve B-cells (CD19+_IgD+_CD27− _{) with}α_IgM/α_{CD40/IL-21 resulted in}

a very small population of B-cells showing proliferation and differentiation into CD38bright_{cells with}

downregulation of surface-IgD but no CD138 upregulation. Under these conditions, a large number of B-cells did not survive (Figure 5A), whereas adding back non-B-cells to the culture showed a survival effect on the naïve B-cells. Stimulation of the sorted naïve B-cells with CpG/IL-2 resulted in some proliferation but no differentiation. Adding back autologous adult non-B-cells to this naïve

B-cell fraction now induced strong proliferation. A CD38bright_{population appeared, although most}

of these cells remained IgD-positive and failed to show any CD138 upregulation. Upon activation the adult naïve B-cells could only produce some IgM upon stimulation with CpG/IL-2, in particular in the presence of non-B-cells, but never any IgG (Figure 5B).

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Figure 4. Age-dependent changes in production and release of immunoglobulins. IgG (A) and IgM (B) levels

were measured in the supernatants by ELISA after 6 days of culture. Plotting multiple experiments per age group (N=3-5). ND means not done.

In contrast, purified adult memory B-cells (CD19+_IgD−_CD27+_{) showed vigorous proliferation}

and differentiation into CD38bright_/CD27high cells with αIgM/αCD40/IL-21, also showing some CD138

expression (Figure 5A) and IgG release (Figure 5B). In the absence of IL-21 though, αIgM/αCD40 did neither result in proliferation nor differentiation, irrespective of the presence of non-B-cells (data not shown). When sorted memory B-cells were activated with CpG/IL-2, almost all B-cells

proliferated and obtained the CD38bright_/CD27high _{phenotype with a subset expressing CD138 and no}

effect of adding non-B-cells (Figure 5A). High amounts of IgG were generated (Figure 5B). Thus, the capacity of B-cells to differentiate into PBs/PCs and to produce IgG was mostly contained in

the memory B-cell pool (IgD –_/CD27+_).

B-cell cultures as a diagnostic tool: proof-of-principle

Many groups have categorized patients with CVID by B cell phenotyping and replication history(14,22), but without extensive exploration at the functional level. We thus investigated in our assays a group of 15 adult CVID patients (Table 1A & supplementary Table 1). Although our test series was small, we could divide our patients into subgroups based on immunophenotyping combined with in-vitro B-cell function.

When compared to healthy adults, B cell stimulation in Group 1 only showed low levels of

proliferation upon stimulation with CpG/IL-2 (Table 1B) or αIgM/αCD40/IL-21 (Table 1C), without

any differentiation to PB/PCs or immunoglobulin production. In Group 2 CpG/IL-2 showed reduced

proliferation and differentiation with low numbers of CD38bright_{PBs but no CD138}+_{PCs, and low levels}

of IgM only. Stimulation with αIgM/αCD40/IL-21 was ineffective. In Group 3, CpG/IL-2 induced

normal proliferation and differentiation into CD38bright_/CD27high _{PBs without CD138}+_{PC phenotype.}

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Figure 5. Activation and reactivity depend on B-cell differentiation. (A) Sorted naïve B-cells (IgD+_/CD27−_{) and}

switched memory B-cells (IgD−_/CD27+_{) were cultured for 6 days as described before, with or without re-addition}

of non B-cells. Representative examples of changes in B-cell phenotype, as well as proliferation into PBs and PCs are shown. (B) Measurement of IgG and IgM levels in the supernatant, after 6 days of culture, by ELISA. (N=3) ND means not done. Stimulation with αIgM/αCD40 gave neither proliferation nor differentiation, irrespective of the presence of T-cells (data not shown).

B cells grow up: Studies of B cell activation, proliferation and differentiation in primary antibody deficiencies - Thesis

UvA-DARE (Digital Academic Repository)