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Original article
Upregulation of RCAN1 causes Down syndrome-like immune dysfunction
  1. Katherine R Martin1,
  2. Daniel Layton2,
  3. Natalie Seach2,
  4. Alicia Corlett1,
  5. Maria Jose Barallobre3,
  6. Maria L Arbonés3,4,
  7. Richard L Boyd2,
  8. Bernadette Scott5,
  9. Melanie A Pritchard1
  1. 1Department Biochemistry and Molecular Biology, Monash University, Clayton, Victoria, Australia
  2. 2MISCL, Monash University, Clayton, Victoria, Australia
  3. 3Center for Genomic Regulation (CRG), UPF, and Centro de Investigaciones Biomédicas en Red de Enfermedades Raras (CIBERER), Barcelona, Spain
  4. 4Institut de Biologia Molecular de Barcelona (IBMB-CSIC), Barcelona, Spain
  5. 5Centre for Functional Genomics and Human Disease, Monash Institute of Medical Research, Monash University, Clayton, Australia
  1. Correspondence to Dr Melanie Pritchard, Department Biochemistry and Molecular Biology, Monash University, Wellington Rd, Clayton, VIC 3800, Australia; melanie.pritchard{at}monash.edu

Abstract

Background People with Down syndrome (DS) are more susceptible to infections and autoimmune disease, but the molecular genetic basis for these immune defects remains undetermined. In this study, we tested whether increased expression of the chromosome 21 gene RCAN1 contributes to immune dysregulation.

Methods We investigated the immune phenotype of a mouse model that overexpresses RCAN1. RCAN1 transgenic (TG) mice exhibit T cell abnormalities that bear a striking similarity to the abnormalities described in individuals with DS.

Results RCAN1-TG mice display T cell developmental defects in the thymus and peripheral immune tissues. Thymic cellularity is reduced by substantial losses of mature CD4 and CD8 thymocytes and medullary epithelium. In peripheral immune organs T lymphocytes are reduced in number and exhibit reduced proliferative capacity and aberrant cytokine production. These T cell defects are stem cell intrinsic in that transfer of wild type bone marrow into RCAN1-TG recipients restored medullary thymic epithelium and T cell numbers in the thymus, spleen and lymph nodes. However, bone marrow transplantation failed to improve T cell function, suggesting an additional role for RCAN1 in the non-haemopoietic compartment.

Conclusions RCAN1 therefore facilitates T cell development and function, and when overexpressed, may contribute to immune dysfunction in DS.

  • Immunology (including allergy)
  • Molecular genetics

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Introduction

Individuals with Down syndrome (DS) have increased susceptibility to infections, particularly respiratory tract infections, with pneumonia being one of the major causes of early death.1 They are also highly susceptible to viral hepatitis and are more likely to develop certain autoimmune disorders, notably thyroiditis, coeliac disease and Type I diabetes.2 ,3 Thus, the immune phenotype in DS is a combination of immunodeficiency and immune dysregulation. The precise cause of immune dysfunction in DS remains unclear, but defective T cell development in the thymus is one contributor.3–5

Mainstream αβ T cell development occurs within the thymus where bone marrow (BM)-derived T cell precursors undergo a complex programme of maturation and differentiation via interactions with specialised resident cells called thymic epithelial cells (TECs).6 The thymic stromal microenvironment consists predominantly of TECs with contributions from fibroblasts, endothelium and haemopoietically derived conventional dendritic cells (cDC) and can be broadly classified into distinct cortical and medullary compartments.7 ,8 Early T cell progenitors (CD4 CD8) enter the thymus at the cortico-medullary junction and migrate through the cortex where they upregulate both CD4 and CD8 expression before the progenitors become restricted to a single CD4 or CD8 lineage. Subsequent migration to medullary regions imposes further selection and developmental events before mature non-self-reactive T cells are exported to the periphery. Thymic stroma cells, such as cortical and medullary epithelial cells (cTEC and mTEC, respectively) regulate thymocyte development by orchestrating functional T cell receptor selection as well as deletion of potential self-reactive clones (reviewed in ref. 6). A bidirectional interplay also exists, in which interactions between thymocytes and stromal elements reciprocally regulate the organisation and development of the thymic microenvironment.9–13

Morphologically, the DS thymus is typically small and exhibits cortical atrophy, a loss of cortico-medullary demarcation5 and lymphopenia due to a defect in the development of thymocytes.3 In the thymus, there are reduced numbers of phenotypically mature T cells that express high levels of the αβTCR-CD3 complex14 and in the periphery, absolute numbers of CD4 T helper cells and CD8 cytotoxic T cells are reduced.4 Functionally, T lymphocytes isolated from individuals with DS exhibit reduced proliferation upon stimulation3 ,15 ,16 and cytokine production is defective; the consensus is that levels of IFN-γ and TNFα are increased, and IL2 is reduced.17–20 While T cell defects have been identified, it remains unclear how they relate to the genomic alterations that occur in DS. Because DS is caused by extra dosage of genes on human chromosome 21, it is possible that dysregulation of a key immune regulator located on chromosome 21 might lead to this spectrum of immune deficits. One potential candidate is Regulator of Calcineurin 1 (RCAN1).

RCAN1 (also known as DSCR1) is overexpressed in DS21–23 and encodes a protein whose best studied function is to modulate the activity of calcineurin.22 ,24–29 Calcineurin is a protein phosphatase that via the dephosphorylation of NFAT links calcium signalling to transcriptional responses in several cell types throughout the body, including T cells.30 NFAT regulates the transcription of a great diversity of target genes, including the proinflammatory genes, IL2, IL4, IFNγ and TNFα.31 Our hypothesis was that RCAN1 modulates immune responses since it regulates calcineurin. Indeed, previous work demonstrated that mice deficient in Rcan1 (RCAN1-KO) displayed normal T cell development and maturation, but exhibited T cell proliferative defects.27 A decrease in IFNγ production was observed indicating that a lack of Rcan1 expression specifically affected the Th1 (T helper type 1) cell population. RCAN1-KO mice, however, do not represent the overexpression of RCAN1 found with trisomy of chromosome 21 and are, therefore, not a clinically relevant model for analysis of immune dysfunction in DS.

We investigated the consequences of increased RCAN1 expression on immune function by generating transgenic mice (RCAN1-TG) that overexpressed the human RCAN1-1 gene under the control of the endogenous isoform 1 promoter.32 RCAN1-TG mice exhibit a small thymus with striking alterations in the thymic stromal microenvironment in addition to T cell developmental defects that result in reduced numbers of CD4 and CD8 thymocytes. In the periphery, the number and functional capacity of mature CD4 and CD8 T cells is reduced. We suggest that RCAN1 is critical for T cell development and function, and show for the first time that overexpression of RCAN1 causes significant T cell immune defects that resemble the immune deficits described in DS.

Methods

Mice

Mice (CBA × C57Bl/6) overexpressing isoform 1 of the human RCAN1 gene (RCAN1-TG) were generated as previously described.32 Sex and age-matched wild type (WT) mice were used as controls. All mice were aged between 8 and 12 weeks unless otherwise stated. Animals were housed in specific pathogen-free conditions with a 12 h light/dark cycle, and had access to food and water ad libitum. All breeding and experiments were conducted in accordance with Monash University Animal Care and Use Committee guidelines.

Quantitative RT-PCR

RNA was extracted using Trizol (Invitrogen, Carlsbad, California, USA) and cDNA generated using Superscript III (Invitrogen, Carlsbad, California, USA) and random hexamers (Promega, Madison, Wisconsin, USA). PCR was used to analyse amplification of either endogenous mouse Rcan1-1, transgenic human RCAN1-1 or Gapdh as a control using the following primer pairs: mouse Rcan1-1 F5′-CCGTGTGGAATTGTCCTTCT; R5′-ACTTCTTTTGCAGGGAAGCA; human RCAN1-1 F5′-TTGGGACTGTCTTGAGAAAACA; R5′-ATCAGTAATATACATGCACAAA; Gapdh F5′-CTGCCACCCAGAAGACTGTGG; R5′-TGGGAGTTGCTGTTGAAGTCG. GelRED (Biotium, Hayward, California, USA) stained DNA fragments were quantified using a Kodak Image Station 4000MMPro and Carestream Molecular Imaging software V.5.0.2.30 (Carestream, Toronto, Canada). The relative expression levels of immune genes was assessed using a Taqman Mouse Immune Array (4367786) and a Real-Time PCR system (ABI7900HT, Applied Biosystems, Foster city, California, USA) on RNA isolated from T cells that were stimulated for 24 h with 10 µg/ml anti-CD3 and 2 µg/ml anti-CD28.

RCAN1 protein analysis in tissues

Tissue snap-frozen in liquid nitrogen was homogenised in tissue lysis buffer (10 mM Tris–HCL, 150 mM NaCl, 0.1% Sodium dodecyl sulphate (SDS), 1% Na deoxycholate, 1% Triton X-100, pH 7.4, supplemented with protease and phosphatase inhibitors). Protein concentration was estimated using the dendritic cell (DC) protein assay kit (BioRad, Hercules, California, USA). Proteins were fractionated on SDS-polyacrylamide gels, transferred onto polyvinylidene fluoride membranes (Immobilon-P, Millipore, Billerica, Massachusetts, USA) and probed with anti-RCAN1 (1:500) (Sigma Aldrich, St Louis, Missouri, USA); anti-tubulin (1:2000) (Chemicon, Temecula, California, USA). The appropriate secondary antibodies were conjugated to horse radish peroxidase (Dako, Denmark, Glostrup). The proteins were visualised using SuperSignal West Pico chemiluminescent substrate (Pierce, Rockford, Illinois, USA) and exposed to Biomax film (Kodak Scientific, Rochester, New York, USA).

Calcineurin activity assay

The Biomol Green Calcineurin Assay kit (Enzo Life Sciences International, Plymouth, Pennsylvania, USA) was used to measure the levels of enzymatic calcineurin activity in the thymus and spleen. A second measure of calcineurin activity was performed as described by Hoeffer and colleagues33 using anticalcineurin (1:2000) (Abcam, Cambridge, Massachusetts, USA) to visualise both full length (60 kDa) calcineurin and the activated cleavage product (48 kDa).

Thymic stromal cell preparation

Thymic stromal cells were isolated from the thymus using a series of enzymatic digestions as previously described.34 Briefly, thymus was digested in Collagenase/DNase with the final digestion performed in Collagenase/Dispase/DNase (Roche, Mannheim, Germany). The single-cell suspensions were passed through 100 μm mesh to remove debris. Excluding the first thymocyte wash, all digestion fractions were pooled and cells counted using a Z2 Coulter Counter (Beckman Coulter, Fullerton, California, USA).

Cell surface marker staining for flow cytometric analysis

Single-cell suspensions of thymus, spleen and LNs were obtained by mechanical digestion in cold Phosphate buffered saline (PBS) supplemented with 0.2% Bovine serum albumin (BSA) (FACS buffer). Single-cell suspensions of freshly dissected BM were obtained by flushing tibias and femurs with cold PBS supplemented with 0.2% BSA with a 26-gauge needle. The marrow was then suspended by gently pipetting through a 22-gauge needle. Cells were recovered by centrifugation at 470×g for 5 min at 4°C. Cell counts were determined by gating viable cells based on cell size using a Z2 Coulter Counter (Beckman Coulter, Fullerton, California, USA). For flow cytometric analysis, 3×106 cells were incubated with the conjugated primary antibodies for 30 min at room temperature, washed with FACS buffer and analysed on a multiparameter FACSCalibur (BD Biosciences, San Diego, California, USA). Data were analysed using GateLogic (Inivai) or FlowJo (Treestar).

For flow cytometric analysis of TECs, 5×106 cells from each thymus digest were stained with appropriate antibodies. Thymic epithelium is classified as negative for haemopoietic marker CD45 and positive for both epithelial cell adhesion molecule (EpCAM) and major histocompatibility (MHC) class II. TECs can be further divided into medullary or cortical epithelium based on positive expression of UEA1 and Ly51, respectively.34 Sample data from CD45EpCAM+ cells were acquired using the BD FACSCanto II flow cytometer (BD Biosciences, San Diego, California, USA) and analysed using FACSDiva software V.6.1.1 (BD Biosciences, San Diego, California, USA).

The following fluorochrome-labelled antibodies against murine antigens were used and were from either BD Biosciences or eBiosciences: FITC and PE-conjugated anti-CD3 (clone 145-2C11); FITC, PE and PercP-conjugated anti-CD4 (clone RM4-5); PE and PercP-conjugated anti-CD8α (clone 53-5.8); allophycocyanin-conjugated anti-TCRβ (clone H57-597); PE-conjugated anti-NK1.1 (clone PK136); PercP-conjugated anti-CD45R/B220 (clone RA3-6B2); FITC or PE-conjugated anti-CD11b (clone M1/70); FITC or PE-conjugated anti-Ly-6G/Ly-6C (Gr-1) (clone RB6-8C5); allophycocyanin-conjugated anti-CD117 (c-kit) (clone 2B8); PE-conjugated anti-Sca-1 (clone D7); FITC or PE-conjugated anti-CD11c (clone HL3); FITC or PE-conjugated CD19 (clone 1D3); allophycocyanin-conjugated anti-CD25 (clone PC61); FITC-conjugated anti-CD44 (clone IM7); allophycocyanin-conjugated anti-IgM (II/41); PE-conjugated anti-CD43 (S7). For analysis of TECs antibodies were from BD Pharmingen (San Diego, California, USA): MTS-15, anti-EpCAM (clone G8.8), anti-Ly51 (clone 6C3), anti-I-A/I-E (clone M5/114.15.2), anti-CD45 (clone 30-F11), anti-CD31 (clone MEC13.3), anti-CD8α (clone 53-6.7), anti-CD4 (clone GK1.5), anti-TCRβ (clone H57-597). The lectin Ulex europaeus agglutinin 1 (UEA-1) was from Vector Laboratories (Burlingame, California, USA). Secondary reagents were Streptavidin antigen-presenting cell (APC) (BD PharMingen, San Diego, California, USA) and antirat IgG2cFITC (Southern Biotech, Birmingham, UK) for detection of MTS-15.

BM transplantation

BM cells were isolated from the tibias and femurs of five female mice per genotype, pooled and depleted of T cells using a Thy1.2 (clone 30-h12) antibody and antimouse IgG Dynal Beads. Mice were irradiated with 12 Gray (Gy), administered in 2 doses of 6 Gy, 3 h apart. After irradiation, 5×106 BM cells were injected into the tail veins of five recipients per group as per table 1. Mice were treated with 0.25 mg/ml enrofloxacin (Baytril 25 oral, Bayer AG, Leverkusen, Germany) for 14 days post-transplant. To allow reconstitution of the immune system mice were left for 14 weeks.

Table 1

The scheme adopted for BM transplantation

T cell proliferation assay

T cells were purified from the spleen using a Pan T cell Isolation kit (Miltenyi Biotech, Bergisch Gladbach, Germany) and an AutoMACS Separator (Miltenyi Biotech, Bergisch Gladbach, Germany) or LNs were excised, dissociated and the single-cell suspensions used to assess proliferation. Cells were plated in Roswell Park Memorial Institute (RPMI) 1640 supplemented with 10% FCS, 100 U/ml penicillin, 200 μg/ml streptomycin, 200 mM L-Glutamine and 0.05 mM 2-mercaptoethanol into either 96 (2×105 cells per well) or 6 (3×106 cells per well) well tissue culture dishes containing either 3 µg/ml or 10 µg/ml of plate bound anti-CD3ɛ (epsilon subunit) and costimulated with 2 µg/ml anti-CD28. For addition of exogenous IL2, cells were seeded into 3 µg/ml anti-CD3ɛ-coated wells and the media supplemented with 50 U per well of IL2 (Roche, Mannheim, Germany). Proliferation was assessed using a CyQUANT Cell Proliferation Assay (Invitrogen, Carlsbad, California, USA).

Cytometric bead assay

Cytokine secretion from activated T cells was investigated using a Th1/Th2 Cytometric Bead Array (#551287, BD Biosciences, San Diego, California, USA). Cytokine release was normalised to the number of viable cells determined at the time of the assay.

Statistical analysis

Statistical analysis was performed using GraphPad Prism V.2.0 software (GraphPad Software, La Jolla, USA). Where there were two groups, a two-tailed non-parametric t test was employed. For analysis of experiments containing three or more groups, a one-way ANOVA and Kruskal Wallis test was used.

Results

RCAN1 is expressed in the thymus, and its upregulation does not inhibit calcineurin activity

We first established that endogenous Rcan1-1 was expressed in the WT murine thymus in both T cells and TECs (figure 1A) and demonstrated expression of the RCAN1-1 transgene in the transgenic thymus (figure 1B (tran),C). In some genetic models upregulation of RCAN1 has been shown to inhibit calcineurin.22 ,26 We measured calcineurin enzymatic activity in whole thymus (figure 1D) and spleen (see online supplementary figure S1) and saw no significant differences in the steady-state levels of calcineurin activity in either organ when RCAN1-TG mice were compared with WT controls. This was confirmed by western blotting using an antibody that detects both full length calcineurin and a constitutively active 48 kDa cleavage fragment.35 ,36 Again, we observed no difference in the amount of the 48 kDa cleavage product between RCAN1-TG and WT mice (figure 1E) confirming that the RCAN1 transgene does not appear to be acting to inhibit steady-state calcineurin activity in either thymus or spleen.

Figure 1

RCAN1 expression and calcineurin activity in the thymus of wild type (WT) and RCAN1-TG mice. (A) Endogenous Rcan1–1–specific RT-PCR of RNA extracted from T cells or thymic epithelial cells derived from three WT thymuses. (B) Quantitative PCR using primers specific to either the endogenous mouse Rcan1-1 gene or to the human transgene (Tran) RCAN1-1 were used on RNA extracted from RCAN1-TG thymuses. Both transcript species were present in the thymuses of transgenic animals. Results were normalised to Gapdh and presented as fold change relative to the endogenous gene. (C) Western blot analysis confirmed up regulation of RCAN1 in the TG thymus. (D) Calcineurin enzymatic activity was measured in the thymus and was no different in RCAN1-TGs compared with controls. (E) Western blotting revealed there was no difference in calcineurin levels between WT and RCAN1-TG mice with respect to both the 60 kDa isoform and a 48 kDa cleavage product that represents a constitutively active form. A representative blot is shown. Intensity of the 48 kDa band is normalised to its respective 60 kDa band. n = 3 mice per group. The data are presented as mean ± SEM. ns, not significant.

Overexpression of RCAN1 leads to reductions in lymphocyte subsets in the thymus, spleen and LNs

To determine the consequences of RCAN1 overexpression on the immune system, we assessed the cellular content and composition of the BM, thymus and peripheral immune organs by flow cytometry. Our results showed comparable numbers of total BM cells isolated from WT and RCAN1-TG mice (figure 2A), as well as similar numbers and proportions of linSca1+ckit+-staining haemopoietic stem cells (HSCs figure 2B,C), granulocytes, macrophages and DCs (see online supplementary figure S2). Taken together, these data indicate that upregulation of RCAN1 does not overtly affect BM cellularity or composition.

Figure 2

T cell development is impaired in RCAN1 transgenic mice. (A) Absolute numbers of cells isolated from the bone marrow (BM), thymus, lymph nodes (LN) and spleen of wild type (WT) and RCAN1-TG animals were counted. A significant reduction in the numbers of cells isolated from the thymus and LN of RCAN1-TG mice was observed. (B) The total number and (C) relative proportion of haemopoietic stem cells was unchanged in RCAN1-TG BM. In the thymus, (D) the total number and (E) relative proportions of T cells in various stages of development were significantly altered by RCAN1 overexpression. In the peripheral immune organs of the LN (F and G) and spleen (H and I) RCAN1 overexpression caused reductions in T cell populations, while B cells remained essentially unchanged. Proportions are relative to total organ cell counts. DN (double negative: CD4CD8); DP (double positive: CD4CD8); B Cell (B220+). Black bars=WT; white bars=RCAN1-TG. n=3–5 mice per genotype. The data are presented as mean±SEM. *p<0.05, **p<0.01, ***p<0.001.

By contrast, there was a significant reduction in the absolute number of cells in the thymus of RCAN1-TG mice compared with WT controls (figure 2A). Reductions were observed across all thymocyte subsets (figure 2D), most notably in the mature CD4 and CD8 single positive (SP) subsets, which were severely reduced in both proportion and overall number (figure 2D,E). Conversely, the proportion of immature double positive (DP; CD4 CD8) cells was increased, while the double negative (DN; CD4CD8) subpopulation remained unchanged (figure 2E). These observations suggest that in the presence of excess RCAN1 the development and maturation of thymocyte subsets is abnormal.

Assessment of the peripheral immune organs revealed a reduction in total cell number in the LN but not in the spleen of RCAN1-TG mice (figure 2A). Nevertheless, a reduction in total CD3 T cell numbers including both CD4 and CD8 SP T cell subsets was observed in both the LN (figure 2F,G) and spleen tissue (figure 2H,I). B220+ B cell numbers were found to be reduced in the transgenic LN but remained unchanged in the transgenic spleen. Together, these results indicate that overexpression of RCAN1 inhibits T cell development in the thymus with a block at the DP stage, and markedly lowers T cell counts in thymus, spleen and LNs.

RCAN1 overexpression alters proportions of thymic cTEC and mTEC and reduces MHCII expression on APCs

The thymic microenvironment is essential for appropriate T cell education and development. We assessed whether upregulation of RCAN1 affected the TEC populations responsible for mediating T cell development and selection by flow cytometric analysis. Our results clearly demonstrate that overexpression of RCAN1 markedly reduced the absolute number of TECs by approximately 90% (figure 3A). Reductions were observed in both mTEC and cTEC subsets (figure 3B) with the mTEC population the more severely affected (figure 3C). The level of MHCII on the surface of TECs (figure 3D), DC, macrophages and B cells (figure 3E–G) within the RCAN1-TG thymus was also reduced.

Figure 3

RCAN1 overexpression alters thymic cTEC and mTEC numbers and reduces major histocompatibility class II (MHCII) expression on thymus antigen-presenting cells. (A) Compared with wild type mice (black bars), the number of TECs was markedly reduced in RCAN1-TG animals (white bars). While the absolute numbers of both mTEC and cTEC populations was decreased in RCAN1-TG mice (B), proportionally, mTECs were significantly decreased and cTECs were significantly increased in RCAN1-TG mice (C). Proportions are relative to total organ cell counts. In RCAN1-TG mice, the level of MHCII expression was significantly reduced for all cell types examined including TECS, DCs, macrophages and B cells (D–G). Percentages on FACS plots represent the proportions of cells within the gates shown. TEC (thymic epithelial cell); mTEC (medullary thymic epithelial cell); cTEC (cortical thymic epithelial cell); DC (dendritic cell); MAC (macrophage). n=5 mice per group. The data are presented as mean±SEM. ***p<0.001.

Overexpression of RCAN1 impairs peripheral T cell proliferation and lowers MHCII on splenic APCs

Rcan1 expression was induced in murine peripheral T cells upon mitogenic stimulation.27 We show that RCAN1 transgene mRNA is similarly induced (see online supplementary figure S3). As thymic T cell development appears to be defective in RCAN1-TG mice, we investigated the proliferative capacity of transgenic T cells derived from either spleen or LN and found that their proliferative capacity was significantly decreased in both organs (figure 4A,B). We assessed the capacity of T cells to secrete the cytokines IL2 and IFNγ and found that secretion was significantly reduced in RCAN1-TG T cells from both organs (figure 4C). Since IL2 is a major proliferative signal for T cells, the proliferative defect observed in RCAN1-TG T cells may be explained by a lack of IL2 in the culture medium. As expected, addition of IL2 to the culture medium induced a significant enhancement in the proliferative capacity of WT T cells, but the same treatment failed to increase the proliferation of RCAN1-TG T cells (figure 4D). To ensure RCAN1-TG cells had the potential to respond to IL2 ligand, we quantified the expression levels of the IL2 receptor and found no significant difference between WT and RCAN1-TG T cells (figure 4E). Taken together, these data indicate that RCAN1-TG T cells are functionally compromised. We next assessed the levels of MHCII expression on antigen-presenting cell populations within the spleen. MHCII expression on DCs and macrophages was significantly lower in RCAN1-TG spleens, but B cell-MHCII expression was unchanged (figure 4F–H).

Figure 4

Overexpression of RCAN1 results in defective T cell proliferation and function in the spleen and lymph node (LN) and reduces major histocompatibility class II (MHCII) expression on splenic antigen-presenting cells. T cells isolated from wild type (WT) (black bars) or RCAN1-TG (white bars) spleens (A) or LN (B) were either left untreated or stimulated with anti-CD3 plus anti-CD28 and their proliferation monitored over 3 days. RCAN1-TG cells had reduced proliferative capacity in response to mitogen stimulation. We quantified cytokines secreted into the media from T cells treated as in (A). The amount of IL2 and IFNγ secreted from RCAN1-TG T cells was significantly reduced (C). T cells isolated from WT and RCAN1-TG animals were treated as described in (A) and the ability of exogenous IL2 to rescue proliferation was determined. Incubation of RCAN1 overexpressing T cells with IL2 did not restore T cell proliferation to the level of WT T cells (D). Isolated T cells treated as above were assayed for IL2R expression. Results were normalised to Gapdh and are presented as fold difference compared with the WT control. IL2R levels were comparable in RCAN1-TG and WT cells (E). In RCAN1-TG mice MHCII expression on splenic dendritic cells (DCs) and MACs was significantly reduced while B cell MHCII expression was unchanged (F–H). n=3 mice per genotype, performed in triplicate. The data are presented as mean±SEM. *p<0.05, **p<0.01.

WT BM restores T cell developmental and TEC defects in RCAN1-TG mice

While thymic epithelium is critical for proper T cell development and function,8 the development and organisation of the thymic stroma is reciprocally dependent on contact with specific thymocyte subsets.9–13 Therefore, it was unclear if the defect in the RCAN1-TG was due to an intrinsic defect in the T cell population, or if the T cell developmental abnormality was a result of a dysfunctional thymic stromal compartment. To resolve this issue, lethally irradiated mice were transplanted with BM isolated from either WT or RCAN1-TG mice according to the scheme shown in table 1. To determine if reconstitution of the immune system by donor BM had occurred after 14 weeks, we isolated splenocytes and performed immunoblotting with an antibody that specifically recognises the RCAN1 transgene (figure 5A). As expected, no transgene expression was detected in WT animals receiving WT BM (WT>WT). However, when WT animals received RCAN1-TG BM (TG>WT), RCAN1 transgene protein was detected, indicating successful reconstitution of the WT immune system with donor TG BM.

Figure 5

Reconstitution of RCAN1-TG mice with wild type (WT) bone marrow (BM) restores mTEC numbers and T cell development in the thymus. Lethally irradiated WT and RCAN1-TG mice were transplanted with BM cells donated from either WT or RCAN1-TG mice. (A) Anti-hRCAN1 immunoblotting on cells isolated from the spleen demonstrated that all BM transplants successfully reconstituted the immune system. The absolute number (B) and proportion (C) of double positive T cells was increased in WT thymuses that received RCAN1-TG BM. WT animals reconstituted with RCAN1-TG BM showed a considerable reduction in the proportion of single positive CD4 (E) and CD8 (G) T cells. These defects were restored to WT levels when RCAN1-TG mice received WT BM. Proportions are relative to total organ cell counts. The presence of WT BM in RCAN1-TG mice resulted in the number of mTECs but not cTECs being restored to WT levels, while the presence of RCAN1-TG BM in WT reduced the mTEC population but had no effect on cTECs (H). n=5 five mice per group. The data are presented as mean±SEM. *p<0.05, **p<0.01, ***p<0.001. mTEC (medullary thymic epithelial cell); cTEC (cortical thymic epithelial cell).

To determine whether the T cell developmental defect observed in RCAN1-TG mice resided in the haemopoietic or non-haemopoietic (stromal) compartment, the BM, thymus, spleen and LN were excised from all groups of chimaeric mice and their cellularity assessed. No significant differences were found in either the number or proportion of HSCs isolated from any of the experimental groups (see online supporting information figure S4). Determinations of absolute T cell numbers (figure 5B,D,F) and proportions (figure 5C,E,G) clearly indicate that transfer of WT BM into TG recipients restored the cellularity of the thymus (compare WT>TG with TG>TG). Interestingly, when WT mice received RCAN1-TG BM (TG>WT) they displayed a small but significant increase in the proportion of DP thymocytes compared with the WT>WT control (figure 5C) which is consistent with our earlier results and seems to indicate that a larger proportion of TG thymocytes remain in the DP state. Concomitantly, these mice displayed a significant decrease in the proportion of SP CD4 and CD8 thymocytes (figure 5E,G, respectively; compare TG>WT with WT>WT). Conversely, if BM from WT mice was transplanted into RCAN1-TG mice (WT>TG) the absolute numbers and proportions of CD4 and CD8 SP thymocytes were restored.

Since distinct thymocyte subsets promote the differentiation of thymic stromal cells,6 ,8 ,12 we assessed the ability of WT BM to restore the thymic stromal cell compartment. Examination of total epithelial cellularity following transfer of WT BM into RCAN1-TG animals (WT>TG) revealed a significant increase in the absolute number of TECs compared with RCAN1-TG mice reconstituted with TG BM (TG>TG) (figure 5H). In particular, mTEC were preferentially restored. Upon transplantation of TG BM, WT recipients exhibited a significant decrease in mTEC number. These results indicate that the T cell development and maturation defect observed in RCAN1-TG mice is intrinsic to the haemopoietic precursor cell population since both thymocytes and stromal cells are restored by WT donor BM.

WT BM restores T cells in RCAN1-TG peripheral immune organs

As reconstitution of RCAN1-TG mice with WT BM restored thymic T cells, we investigated if this would be reflected in the peripheral immune organs. Transplantation of WT BM into transgenic recipients (WT>TG) significantly increased both the number and proportion of CD4 and CD8 T cell subsets (figure 6A–D). As expected, transplantation of RCAN1-TG BM into WT recipients (TG>WT) resulted in deficient numbers of both CD4 and CD8 SP T cells in the spleen (figure 6A–D). These findings were reproduced upon analysis of the LN (figure 6E–H).

Figure 6

Transplantation of wild type (WT) bone marrow (BM) into RCAN1-TG mice restores normal single positive T cell numbers in the spleen and lymph node (LN) but was unable to restore T cell function or major histocompatibility class II (MHCII) expression on antigen-presenting cells. The presence of WT BM in RCAN1-TG mice restored CD4 (A and B) and CD8 (C and D) T cells to WT levels in the spleen. Similar analysis of the LN also revealed increases in the number and proportion of CD4 (E and F) and CD8 (G and H) T cells in RCAN1-TG mice transplanted with WT BM. Proportions are relative to total organ cell counts. T cells isolated from LN of chimaeric mice were stimulated with anti-CD3 plus anti-CD28 and their proliferation measured after 48 h. Reconstitution of RCAN1-TG mice with WT BM did not restore T cell proliferation to WT levels (I). Secretion of IFNγ (J) and IL2 (K) remained low in RCAN1-TG mice upon reconstitution with either WT or TG BM. Conversely, the presence of RCAN1-TG BM did not adversely affect cytokine secretion in WT mice. Dendritic cells, MACs and B cells were isolated from the spleens of chimaeric mice and the level of cell surface MHCII expression analysed. In RCAN1-TG recipients the level of MHCII expression was significantly reduced for all cell types examined (l–n). Notably, the presence of WT BM did not rescue this phenotype. n=5 mice per group. The data are presented as mean±SEM. *p<0.05, **p<0.01, ***p<0.001.

WT BM was unable to restore normal T cell function in RCAN1-TG mice

To determine if T cell function was restored, LN-derived T cells were assessed for their proliferative capacity and ability to release the cytokines IL2 and IFNγ. Consistent with our earlier findings, T cells isolated from WT>WT control mice proliferated significantly more than those derived from TG>TG mice (figure 6I). Interestingly, the proliferative capacity of peripheral T cells was not restored in WT>TG mice and proliferative function was not reduced in TG>WT (figure 6I). Consistent with these findings, secretion of the Th1 cytokines IL2 and IFNγ, was not rescued in WT>TG, whereas in TG>WT IL2 and IFNγ secretion remained at WT levels (figure 6J,K). Finally, in groups where cytokine production remained deficient, MHCII expression on spleen-derived APCs was significantly lower (figure 6L–N).

Discussion

In this study using genetically modified mice as a disease model, we examined whether overexpression of RCAN1 had the potential to contribute to the T cell deficits associated with DS. Overexpression of RCAN1 in mice resulted in immune defects equivalent to those observed in DS; namely impaired T cell development and maturation, altered thymic microenvironment and T lymphocyte functional deficiencies that affect both proliferation and cytokine production. Our study found that T cell developmental defects were intrinsic to the stem cell compartment as transfer of WT BM restored the thymic medullary epithelium and T cell numbers in thymus, spleen and LNs of TG mice. However, BM transplantation failed to improve peripheral T cell function which suggests that RCAN1 has an additional but hitherto undefined role in the non-haemopoietic stromal compartment.

It has been established that RCAN1 is able to modulate (either facilitate or inhibit) calcineurin activity.22 ,24–29 In our study, using two independent methods of measuring calcineurin activity, we were unable to show that overexpression of RCAN1 had any measurable effect on calcineurin activity. These results are consistent with calcineurin activity determinations in other tissues of RCAN1-TG mice, including the brain.37 Therefore, the immune abnormalities observed in our RCAN1-TG mice are probably independent of the calcineurin pathway.

Since the immunodeficiencies in DS were replicated in our RCAN1-TG mice, upregulation of RCAN1 appears responsible, at least in part, for the immune deficiencies associated with DS. Both humans with DS5 and RCAN1-TG mice display thymic defects including reduced size of the thymus, abnormal T cell development and lymphopenia. In both, there is a reduction in the number of CD4 and CD8 T cell subsets in the periphery, and also these T cells display functional deficits including reduced proliferative capacity and decreased production of IL2. However, overexpression of RCAN1 in mice does not entirely simulate the T cell phenotype of DS. For example, mitogenic stimulation of T cells in RCAN1-TG mice resulted in the secretion of lower levels of IFNγ compared with controls (figure 4C), while similarly treated T cells isolated from individuals with DS secrete increased levels.5 Given that chromosome 21 is estimated to contain between 300 and 400 genes, many of which are relevant to immune system function, upregulation of any single gene would not entirely recapitulate the immune dysfunction observed in DS. Thus RCAN1 likely acts in combination with other chromosome 21 genes to generate the spectrum of immune system deficits of DS. In this regard, it should be noted that people with DS have a marked B cell cytopenia, and skewing of their B cell populations suggests that maturation of B cells is also defective.38 Immunoglobulin levels in DS are abnormal and antibody responses to a variety of antigens are low.3 ,38 In this study, we have not explored the consequences of RCAN1 overexpression on the B cell compartment.

A combination of export of T cells from the thymus and their proliferation in the periphery contribute to the establishment of the T cell pool.39 We found that overexpression of RCAN1 resulted in a marked reduction of SP T cells in the thymus, spleen and LNs. These observations can be explained by our findings that excess RCAN1 inhibited the maturation and differentiation of T cells, consistent with a T cell progenitor block at the DP stage, and caused a reduction in the proliferative capacity of peripheral T cells. We have not directly tested thymocyte export. Notably, a recent report found that decreased thymic output accounted for diminished numbers of T cells in children with DS rather than a reduced capacity of the T cells to proliferate.40 However, their ex vivo proliferation data is not shown.

The loss of medullary epithelium in the thymus of TG mice was also consistent with the observed decrease in mature SP thymocyte subsets, since final selection/development events occur in the thymic medulla before the export of mature T cells subsets to the periphery. The concomitant loss of mTECs in mouse models of defective T cell development due to lack of ‘cross-talk’ between mature thymocyte subsets and thymic stroma cells is well documented.11 ,12 Interestingly, we also observed a significant decrease in the level of MHCII expression on TECs from RCAN1-TG mice. Given that a threshold level of MHCII expression is required for normal thymocyte maturation, in particular for the maturation of CD4 SP thymocytes,41 and lower levels have been associated with defects in T cell development,41–43 reduced MHCII expression in the RCAN1-TG thymus could contribute to the observed thymic T cell developmental deficits. Similarly, high MHCII expression on peripheral APCs is essential for T cell activation and function.41 ,44 ,45 Thus, low levels of APC-associated MHCII in peripheral immune organs of RCAN1-TG mice would prevent optimal T cell function and contribute to their reduced proliferative capacity upon TCR ligation. How and why an increased level of RCAN1 affects MHCII expression is unknown.

When the immune system of RCAN1-TG mice was reconstituted with WT BM, the T cell numbers and defects associated with the thymic microenvironment were restored. This may not be surprising since donor-derived TECs have been found to repopulate the thymus of the recipient indicating that BM cells contain the precursors of functional TECs.46 Conversely, when WT mice received RCAN1-TG BM they acquired the same T cell developmental and structural thymic defects that are observed in RCAN1-TG mice. These data strongly indicated a defect intrinsic to the haemopoietic compartment of RCAN1-TG mice which impedes proper T cell development and maturation. However, T cells isolated from RCAN1-TG mice repopulated with WT BM were still deficient in proliferative capacity and cytokine production. This indicates a role (at least in part) for the RCAN1-TG stromal microenvironment in the maintenance of T cell activation and function. Interestingly, this reduction in peripheral T cell function was associated with reduced MHCII expression on APCs. Thus, it appears that despite transplantation with WT BM, APC development (as determined by reduced MHCII expression) was defective in immune organs where the stromal compartment remained RCAN1-TG-derived. Conversely, when WT mice were transplanted with BM from TG mice, fewer mature T cells developed and migrated to the periphery as a result of intrinsic RCAN1 expression, yet their function was restored, perhaps because APC-MHCII expression in the periphery was normal.

In conclusion, we reveal a link between overexpression of a DS-related gene RCAN1 and T cell deficiency. Our model could well be added to the growing list of diseases with combined features of immunodeficiency and autoimmunity described by Mackay and colleagues.47 However, the molecular pathways that mediate the immunosuppressive effects of excess RCAN1 are as yet unidentified. Further knowledge should not only allow for new therapies to ameliorate the immune deficits of DS, but could also be applicable to treatment of autoimmune disease and impending graft rejection after organ transplant. In fact, a RCAN1 antagonist has already been shown to decrease the expression of RCAN148 and, thus, could improve immune function in DS, and a RCAN1 agonist could suppress the immune system as a therapy for autoimmune disease or a failing tissue transplant. Our RCAN1-TG model could well be useful for assessing effects of RCAN1 antagonists or agonists.

Acknowledgments

We would like to thank Professor Ian Mackay and Dr Georg Bohn for critical reading of the manuscript. This work was supported by the Fondation Jérôme Lejeune; and philanthropic grants from the Judith Jane Mason & Harold Stannett Williams Memorial Foundation managed by ANZ Trustees; the APEX Foundation for Research into Intellectual Disability; the CASS Foundation; and the LEW Carty Charitable Fund. MJB is supported by the CIBERER. The Authors have no conflicting financial or other competing interests in relation to this work.

References

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Footnotes

  • Contributors KRM: conception and design, acquisition, analysis and interpretation of data; drafting the article; critical revision; final approval. DL: conception and design, acquisition, analysis and interpretation of data; drafting the article; critical revision; final approval. NS: conception and design, acquisition, analysis and interpretation of data; drafting the article; critical revision; final approval. AC: acquisition, analysis and interpretation of data; drafting the article; critical revision; final approval. MJB: acquisition, analysis and interpretation of one aspect of the data; final approval. MLA: analysis and interpretation of one aspect of the data; critical revision; final approval. RLB: interpretation of TEC data; final approval. BS: conception and design, interpretation and critique of the data, drafting the article; critical revision; final approval. MAP: conception and design, analysis and interpretation of data; drafting and final writing of the article the article; critical revision; final approval, submission.

  • Competing interests None.

  • Provenance and peer review Not commissioned; externally peer reviewed.

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