The Size of the Plasmacytoid Dendritic Cell Compartment Is a Multigenic Trait Dominated by a Locus on Mouse Chromosome 7

Adam-Nicolas Pelletier, Fanny Guimont-Desrochers, Michelle P. Ashton, Thomas C. Brodnicki and Sylvie Lesage
J Immunol June 1, 2012, 188 (11) 5561-5570; DOI: https://doi.org/10.4049/jimmunol.1102136

Abstract

Plasmacytoid dendritic cells (pDC) compose one of the many distinct dendritic cell subsets. The primary function of pDC is to potently produce type 1 IFNs upon stimulation, which is highly relevant in antiviral responses. Consequently, the ability to manipulate the size of the pDC compartment in vivo may increase the capacity to clear viral infections. In an attempt to identify genetic loci affecting the size of the pDC compartment, defined by both the proportion and absolute number of pDC, we undertook an unbiased genetic approach. Linkage analysis using inbred mouse strains identified a locus on chromosome 7 (Pdcc1) significantly linked to both the proportion and the absolute number of pDC in the spleen. Moreover, loci on either chromosome 11 (Pdcc2) or 9 (Pdcc3) modified the effect of Pdcc1 on chromosome 7 for the proportion and absolute number of pDC, respectively. Further analysis using mice congenic for chromosome 7 confirmed Pdcc1, demonstrating that variation within this genetic interval can regulate the size of the pDC compartment. Finally, mixed bone marrow chimera experiments showed that both the proportion and the absolute number of pDC are regulated by cell-intrinsic hematopoietic factors. Our findings highlight the multigenic regulation of the size of the pDC compartment and will facilitate the identification of genes linked to this trait.

The primary characteristic defining plasmacytoid dendritic cells (pDC) is their ability to potently produce type 1 IFNs upon stimulation, conferring them with a key antiviral role (1–3). Indeed, mice presenting with a higher proportion of pDC produce more IFN-α in response to hypomethylated CpG (4). The definitive in vivo contribution of pDC toward viral clearance was recently assessed by infecting pDC-depleted mice with either murine CMV or vesicular stomatitis virus (5). In the absence of pDC, murine CMV-infected mice produced less type 1 IFN and presented with an altered NK cell response, whereas vesicular stomatitis virus-infected mice showed a significant decrease in the survival and recruitment of CD8 T cells. These observations demonstrate that in the context of specific viral infections, pDC contribute to both innate and adaptive immunity. Therefore, understanding the regulation of the pDC compartment may allow for the manipulation of pDC number and/or proportion, which should enhance immune responses toward viral infections.

pDC are present at a low proportion in secondary lymphoid tissues and compose ∼1% of total spleen cells. The size of the pDC compartment is determined, as for any other hematopoietic subset, by the homeostatic pressures conferred to this cellular subset. As for other dendritic cell (DC) subsets, pDC are derived from Flt3+ precursors in the bone marrow (6, 7). However, pDC are relatively quiescent, suggested by the low rate of incorporation of the DNA base analog BrdU (8–10). In addition, parabiotic experiments have demonstrated that the half-life of pDC is notably shorter than that of other DC subsets (9). Hence, the quiescence and short half-life of pDC may contribute toward defining their small compartment size (3).

Molecular determinants favoring pDC differentiation have also been identified. A cDNA subtraction library demonstrated that Spi-B was preferentially expressed in human pDC relative to monocyte-derived DCs (11). Overexpression and knockdown experiments conclusively demonstrate that Spi-B is a determining transcription factor promoting the differentiation of human pDC in vitro (11, 12). IFN regulatory factor (IRF)-8 also proved to be a key transcription factor in pDC differentiation and function, as Irf8-deficient mice presented with a much reduced proportion of pDC along with a deficiency in type I IFN production upon viral infection (13). More recent studies have proposed that the E2-2 transcription factor serves as a specific regulator of the pDC transcription profile, acting upstream of both Spi-B and IRF-8, and revealed its importance in the maintenance of the pDC fate (14, 15). Therefore, the differentiation of pDC is dependent on the timely expression of specific transcription factors.

Although the molecules and pathways relevant to the differentiation of pDC have been identified, the molecular determinants defining the size of the pDC compartment remain to be elucidated. Thus, we implemented an unbiased genetic approach to identify key genetic determinants that regulate the size of the pDC compartment. Asselin-Paturel et al. (4) have previously documented differences in pDC proportion among different inbred mouse strains. Based on these observations, we undertook a linkage analysis of (NOD×B6)F2 mice deficient for Rag1 to identify loci that are linked to the size of the pDC compartment, which is defined by both the proportion and the absolute number of pDC. We found a significant linkage to both traits (proportion and absolute number of pDC) on the proximal end of mouse chromosome 7 and named the locus pDC compartment 1 (Pdcc1). Moreover, we identified genetic interactions between Pdcc1 and a locus on chromosome 11 contributing to the proportion of pDC, named Pdcc2, and between Pdcc1 and a locus on chromosome 9 contributing to the absolute number of pDC, named Pdcc3. These data demonstrated that the size of the pDC compartment in the spleen is a multigenic trait. We further validated the linkage of chromosome 7 to both the absolute number and the proportion of pDC using a congenic mouse strain and demonstrated that the size of the pDC compartment is determined by cell-intrinsic hematopoietic factors.

Materials and Methods

Mice

C57BL/6 (hereafter denoted B6), C3H, BALB/c, NOD, NOR, NZW, B10.Br, B6.Rag1−/− and NOD.Rag1−/− mice were purchased from The Jackson Laboratory and subsequently maintained at the Maisonneuve-Rosemont Hospital animal housing facility (Montreal, QC, Canada). NOD.H2k (16), F1 (B10.Br × NOD.H2k), F1.Rag (B6 Rag1−/− × NOD Rag1−/−), and F2.Rag (F1.Rag−/− × F1.Rag−/−) mice were maintained at the Maisonneuve-Rosemont Hospital animal housing facility. For experiments using NOD.NZW-Chr7 mice, NOD and NZW (both purchased from The Jackson Laboratory) as well as NOD.NZW-Chr7 (17) mice were bred and maintained at St. Vincent’s Institute animal facility (Melbourne, VIC, Australia). Six- to 8-wk-old mice were used for phenotypic analyses. For generating bone marrow chimeras, F1 (B10.Br × NOD.H2k) recipient mice were irradiated at 11 Grays and left to reconstitute for 8 wk with 2 × 106 bone marrow cells from B10.Br and NOD.H2k mice at a 1:1 ratio. The Maisonneuve-Rosemont Hospital ethics committee, overseen by the Canadian Council for Animal Protection, and the animal ethics committee at St. Vincent’s Hospital approved the experimental procedures.

Flow cytometry

Spleens, lymph nodes, and thymus are treated with collagenase (1 mg/ml; type V from Clostridium histolyticum; Sigma-Aldrich) for 15 min at 37°C. Bone marrow is flushed from the femur and tibias. Livers were perfused with PBS and subsequently with collagenase. Livers were then harvested and digested for 25 min at 37°C with 5 ml collagenase. Lungs were perfused with PBS, harvested, and digested for 30 min at 37°C with 1 ml collagenase. All organs were passed through a 70-μm cell strainer (BD Biosciences) to yield single-cell suspensions prior to staining with Abs. CD11c PE-Cy7, CD11c FITC, Siglec-H PE, and B220 allophycocyanin Abs were purchased from BioLegend, mouse pDC Ag-1 allophycocyanin from Miltenyi Biotec, and B220 PerCP from BD Biosciences. All samples were run using an FACSCalibur (BD Biosciences), FACSCanto I (BD Biosciences), or BD LSRII (BD Biosciences) and analyzed using the FlowJo software (Tree Star, Ashland, OR). Note that we implemented a precise gating strategy to allow for the accurate identification of pDC proportion in Rag-deficient mice. Briefly, to remove autofluorescent cells (18), two dump channels are used on the 488-nm laser, namely FL1 and FL2. A live gate is then applied, and doublets are removed. pDCs are subsequently selected based on the CD11clowBst-2+ phenotype. Finally, a strict size exclusion gate is applied to the forward light scatter/side scatter profile, corresponding to the pDC subset.

Linkage analysis

Genomic DNA was isolated from the tails of F2.Rag male and female mice by using the DNeasy blood and tissue kit from Qiagen. Single nucleotide polymorphisms (SNPs) were then detected from the F2.Rag mice DNA using the Illumina mouse low-density linkage panel serviced through The Centre for Applied Genomics at the Hospital for Sick Children. Additional genetic markers were used to delimit the interval on mouse chromosome 7, namely D7Mit178 (3.46 Mb), D7Mit306 (10.99 Mb), and D7Mit191 (19.47 Mb). The logarithm of odds (LOD) scores were obtained through the single- or two-dimensional quantitative trait locus model using the R/qtl package (19) for the R software (version 2.11.1). LOD scores >3.1 were considered significant for single-dimensional analysis according to permutation tests (n = 10,000; p = 0.05), and LOD scores between 2 and 3.1 were considered suggestive. The two-dimensional model required a LOD score of 9.3 for significance, calculated by permutation testing (n = 1000; p = 0.05).

IFN-α production

pDC were sorted from pooled spleens of B6, NOD, NZW, and NOD.NZW-Chr7 mice. Purity was always >95%. A total of 1 × 105 sorted pDC were stimulated for 21 h in vitro using 5 μg/ml CpG ODN 2216 (InvivoGen). The concentration of IFN-α present in the supernatant was assessed with FlowCytomix Mouse IFN-α detection kit (eBioscience).

Measure of proliferation and apoptosis

B10.Br and NOD.H2k mice were sacrificed 24 h after i.p. injection with 1 mg BrdU. Extracellular staining for CD11c and Bst-2 on total spleen cells was performed prior to intracellular staining for BrdU, active caspase-3, and Bcl-2 (BD Biosciences), which were performed using the permeabilization kit BD Cytofix/CytoPerm (BD Biosciences). For BrdU, a 1-h OmniPur DNase I treatment (Calbiochem) was also performed. An isotype Ab control was used for Bcl-2 staining (BD Biosciences), whereas the other intracellular stainings were compared using the fluorescence minus one approach (20).

Statistical analysis

Data were tested for significance using a nonparametric Mann–Whitney U test with a minimal threshold of 0.05. Estimation of the interval coordinates was obtained using a 95% Bayes interval test. All statistical analyses and the F2 distribution were obtained using SPSS 17.0 software.

Results

To identify pDC in the spleen, we took advantage of their CD11clowBst-2+-specific phenotype, in which Bst-2 is detected using the mouse pDC Ag-1 mAb (Fig. 1A) (3). Notably, all CD11clowBst-2+ cells express B220 (Supplemental Fig. 1A), validating the specificity of the CD11c and Bst-2 marker combination in delineating the pDC population. Using these cell-surface markers, Asselin-Paturel et al. (4) documented variation for the proportion of pDC among a panel of inbred mouse strains, in which B6 exhibited the lowest and 129Sv the highest pDC proportion in the spleen. To confirm and extend this study, we measured the proportion of pDC in spleens obtained from five inbred mouse strains, two of which had not been previously documented, namely NOD and NOR strains. In addition, we documented the absolute number of pDC in the spleen of each strain. Similar to that observed by Asselin-Paturel et al. (4), B6 mice exhibited the lowest proportion and number of pDC (Fig. 1B). C3H and BALB/c mouse strains presented slightly increased levels, whereas NOD and NOR mouse strains exhibited the highest proportion and absolute number of pDC in this collection of mouse strains (Fig. 1B).

FIGURE 1.
FIGURE 1.

Inbred mouse strains exhibit variation for the proportion and absolute number of pDC. (A) Representative flow cytometry profiles of CD11c and Bst-2 expression on total spleen cells from a B6 mouse. Application of the live gate (left panel). The percentage of live CD11clow Bst-2+ cells is shown (right panel). (B) pDC proportion (left panel) and absolute numbers (right panel) are indicated for various strains of mice. Each dot represents data for an individual mouse. Data were acquired using a BD FACS Calibur flow cytometer (BD Biosciences). **p < 0.01.

To investigate the genetic basis of pDC variation in mice, we performed a linkage analysis using B6 and NOD mouse strains for the following reasons. First, these two mouse strains, respectively, exhibited the lowest and highest levels of pDC (Fig. 1B), suggesting that these distinct genetic backgrounds provide the greatest chance of detecting genetic linkage to this pDC trait in a (NOD×B6)F2 cohort. Second, the NOD mouse strain is a widely used model for genetic studies of type 1 diabetes (T1D). A number of congenic NOD strains have been generated to confirm T1D loci (21, 22) and, accordingly, could be used to confirm overlapping loci identified in our pDC linkage analysis.

One potential limitation for a linkage analysis of pDC is the sensitivity for measuring variation of pDC levels in a (NOD×B6)F2 outcross. We first validated that CD11clowBst-2+ cells efficiently delineated pDC in both parental strains, namely B6 and NOD mice. We found that Bst-2 and Siglec-H were effectively coexpressed on CD11clow cells from both mouse strains (Supplemental Fig. 1B) in steady-state conditions (23). Hence, we continued using only CD11c and Bst-2 to identify pDC. The difference between B6 and NOD mouse strains for the proportion of pDC was <1%, whereas B6 and NOD mouse strains, respectively, had an average of 0.6 and 1.5% pDC in the spleen (Fig. 1B). To enhance the detection of pDC differences, we postulated that the proportion of pDC would be increased in mice harboring a null mutation for Rag due to the lack of lymphocytes in these mice. Accordingly, we determined the proportion of pDC in B6.Rag1−/− and NOD.Rag1−/− mice, hereafter, respectively, referred to as B6.Rag and NOD.Rag. As Rag-deficient mice lack T and B cells, the gating strategy was adapted to account for a much higher percentage of autofluorescent cells (18) (Fig. 2A; see also Materials and Methods). We found that a larger difference could be detected for the proportion of pDC between NOD.Rag and B6.Rag mice (Fig. 2B) compared with the difference observed between NOD and B6 wild-type mice (Fig. 1B). Indeed, the Rag-deficient background presented a difference of 8% in pDC proportion (an average of 8 versus 16% of pDC was detected in B6.Rag and NOD.Rag mice, respectively), as opposed to a difference of <1% in the Rag-sufficient background. Moreover, the sensitivity in pDC detection was increased upon Rag deficiency, in which, for instance, B6.Rag mice exhibited 8% pDC compared with only 0.6% in B6 mice. Together, these data demonstrate that the sensitivity in detecting pDC is increased in Rag-deficient mice. Correspondingly, as for Rag-sufficient mice, the absolute number of pDC in the spleen was lower in B6.Rag mice relative to NOD.Rag mice (Fig. 2C). As NOD.Rag mice do not develop autoimmunity due to the absence of lymphocytes, these observations further indicate that the increased pDC proportion observed in the NOD mice, relative to the B6 mice, is not a consequence of an underlying autoimmune pathology.

FIGURE 2.
FIGURE 2.

Increased sensitivity in the detection of pDC variation between B6.Rag and NOD.Rag mice. (A) The stepwise logical gating strategy is shown for the detection of pDC in Rag-deficient mice. Representative plots of the analysis of a B6.Rag mouse are shown. 1, Two dump channels on the 488-nm laser are used to remove autofluorescent cells. 2, Application of live gate (doublets are also removed, not shown). 3, pDC are subsequently selected based on the CD11clowBst-2+ phenotype. 4, A strict size-exclusion gate is applied to the forward light scatter (FSC)/side scatter (SSC) profile, corresponding to the pDC subset. The proportion (B) and absolute number (C) of pDC in the spleen of B6.Rag and NOD.Rag mice is shown. Each dot represents data for an individual mouse. Data were acquired using a BD FACS Canto flow cytometer (BD Biosciences). *p < 0.05.

Based on the increased sensitivity for detecting pDC in Rag1-deficient mice and the consequential increased difference in pDC proportion between B6.Rag and NOD.Rag mice, we generated a cohort of 181 (NOD.Rag × B6.Rag)F2 mice (referred to as F2.Rag). The proportion of pDC in F2.Rag mice varied from 1.3 to 39% and showed a normal distribution (Fig. 3A). Genomewide SNP genotyping was performed on this F2.Rag cohort by taking advantage of the Illumina low-density platform. The linkage analysis for pDC proportion demonstrated a significant linkage on chromosome 7 with a LOD score of 5.46, well above the significance threshold of 3.1 determined by permutation testing (Fig. 3B). In addition, linkage above the suggestive threshold was observed on both chromosomes 8 and 13, with a locus on chromosome 1 almost reaching the suggestive threshold (Fig. 3B).

FIGURE 3.
FIGURE 3.

Linkage analysis of pDC proportion and absolute number. The distribution of the F2.Rag cohort relative to the proportion (A) and absolute number (C) of pDC are shown. Genomewide LOD score plot (R/Qtl) for the proportion (B) and absolute number (D) of pDC in the F2.Rag cohorts are shown. The dashed and dotted lines, respectively, indicate the significance threshold of p < 0.05 and the suggestive threshold.

The size of the pDC compartment is defined by both the proportion and absolute number of pDC. Therefore, to fully characterize the genetic loci defining the size of the pDC compartment, we performed a linkage analysis on the absolute number of pDC in the spleen of F2.Rag mice. The absolute number of pDC in the F2.Rag cohort exhibited a normal distribution (Fig. 3C) and was significantly linked to the same locus as pDC proportion on mouse chromosome 7 (Fig. 3D). Linkage above the suggestive threshold on chromosomes 1 and 16 was observed, yet chromosomes 8 and 13 were not linked to the absolute number of pDC. Together, these data show that a dominant locus on chromosome 7 is significantly linked to both the proportion and absolute number of pDC, whereas additional suggestive linkages are distinctly linked to either trait.

To delimit the interval associated with the size of the pDC compartment on mouse chromosome 7, the F2.Rag cohort was genotyped for three additional genetic markers (D7Mit191, D7Mit306, and D7Mit178). These three markers are proximal to the SNP rs3720735, the most proximal informative SNP from the Illumina low-density platform. Higher-resolution maps of chromosome 7 for linkage to both the proportion and the absolute number of pDC further delimited the locus, which we have named Pdcc1. Pdcc1 is located at the proximal end of the chromosome 7 with a peak at 47.7 Mb, corresponding to the SNP rs3670807 (Fig. 4A, 4B). The 95% Bayes interval test delimits the Pdcc1 locus between 23.5 and 63.5 Mb. To evaluate the direct relationship of Pdcc1 to the size of the pDC compartment, the F2.Rag mice were segregated according to their genotype at SNP rs3670807 (Fig. 4C, 4D). The F2.Rag mice harboring the NOD/NOD homozygous genotype at the SNP rs3670807 exhibited a higher proportion and absolute number of pDC relative to those mice harboring the heterozygous B6/NOD genotype, which exhibited higher levels of pDC compared with mice harboring the homozygous B6/B6 genotype.

FIGURE 4.
FIGURE 4.

pDC proportion and absolute number are affected by Pdcc1, a dominant locus on chromosome 7. High-resolution maps of the linkage of chromosome 7 to pDC proportion (A) and absolute number (B). The dashed and dotted lines, respectively, indicate the significance threshold of p < 0.05 and the suggestive threshold. Representation of pDC proportion (C) and absolute number (D) in F2.Rag mice segregated according to the genotype for the SNP rs3670807 on chromosome 7. Each dot represents data for an individual mouse. *p < 0.05, **p < 0.01, ***p < 0.001, ****p < 0.0001. B/B mice, Homozygous for B6 alleles; B/N, heterozygous for B6 and NOD alleles; N/N, homozygous for NOD alleles.

The normal distribution of the proportion and absolute number of pDC in F2.Rag mice suggested a multigenic trait, yet only the proximal region of chromosome 7 was significantly associated with this trait. Joint LOD score analyses were subsequently performed to reveal genetic interactions that may further define the loci affecting the proportion of pDC (24). Although suggestive linkages were identified on chromosomes 8 and 13 for the proportion of pDC (Fig. 3B) and on chromosomes 1 and 16 for the absolute number of pDC (Fig. 3D), the joint LOD score analysis indicated that these linked regions were unlikely to interact with one another or Pdcc1 to affect either the proportion or the absolute number of pDC (Fig. 5A, 5B). In contrast, the joint LOD score analysis detected significant genetic interactions between loci on chromosome 7 and 11 that linked to the proportion of pDC and between loci on chromosome 7 and 9 that linked to the absolute number of pDC (Fig. 5A, 5B). Further segregation analysis indicated that these interactions are complex. The two SNPs defining the peak joint LOD score (Chr7, rs3670807; Chr11, rs1341088) were used to segregate mice and determine the mean proportion of pDC for each genotype combination (Fig. 5C). Mice that were NOD homozygous for both SNPs exhibited the highest proportion of pDC. Intriguingly, mice that were B6 homozygous for both SNPs did not exhibit the lowest pDC proportion. The lowest proportion was found in mice that were B6 homozygous for the chromosome 7 SNP, but NOD homozygous for the chromosome 11 SNP. Mice with the remaining genotype combinations had pDC levels that were intermediate or similar to these other genotypes (Fig. 5C). Similarly, segregation of F2.Rag mice according to their genotype at the two SNPs defining the peak joint LOD score (Chr7, rs3670807; Chr9, rs3665206) for the absolute number of pDC also demonstrated a complex interaction (Fig. 5D). The lowest number of pDC was found in mice that were B6 homozygous for the chromosome 7 SNP and NOD homozygous for the chromosome 9 SNP (Fig. 5D). These results reinforced the finding that Pdcc1 is a key genetic determinant for this trait, but also pointed to two distinct loci, namely Pdcc2, located on chromosome 11, and Pdcc3, located on chromosome 9, that can modify the effect of Pdcc1 depending on the genotype combination.

FIGURE 5.
FIGURE 5.

Genetic interactions between loci affect the size of the pDC compartment. (A) Genomewide two-dimensional LOD score analysis of F2.Rag mice for pDC proportion; LOD score = 9.53 for the interaction between chromosomes 7 and 11. (B) Genomewide two-dimensional LOD score analysis of F2.Rag mice for pDC absolute number; LOD score = 9.60 for the interaction between chromosomes 7 and 9. (C) Representation of pDC proportion in F2.Rag mice segregated according to the combined chromosome 7 (rs3670807) and 11 (rs13481088) genotypes. (D) Representation of pDC number in F2.Rag mice segregated according to the combined chromosome 7 (rs3670807) and 9 (rs3665206) genotypes. *p < 0.05. B/B mice, Homozygous for B6 alleles; B/N, heterozygous for B6 and NOD alleles; N/N, homozygous for NOD alleles.

To confirm that Pdcc1 is a key locus in determining the size of the pDC compartment, we took advantage of a congenic NOD strain (NOD.NZW-Chr7) that harbors an NZW-derived chromosome 7 interval (17) (Fig. 6A). Notably, NOD.NZW-Chr7 mice exhibited a significant reduction in both the proportion and absolute number of pDC in the spleen compared with NOD mice (Fig. 6B). Also, there is a tendency toward a reduction in pDC number in all organs examined (Supplemental Fig. 2), suggesting that the decrease in number observed in the spleen is not due to preferential accumulation of pDC in other lymphoid organs in the NOD.NZW-Chr7 congenic mouse. Therefore, changing the genetic interval on chromosome 7 in NOD mice alters the size of the pDC compartment in Rag-sufficient mice. This observation not only confirms our linkage of Pdcc1 to this trait, but also it agrees with the premise that the size of the pDC compartment in both Rag-deficient and -sufficient mice is defined at least in part by similar genetic variants.

FIGURE 6.
FIGURE 6.

Validation of the contribution of chromosome 7 in defining the size of the pDC compartment. (A) Schematic diagram of chromosome 7 for the NOD.NZW-Chr7 mouse strain. Marker placement is shown in Mb according to the National Center for Biotechnology Information m37 build. Pdcc1 is depicted according to the Bayes interval. (B) pDC proportion (top panel) and absolute number (bottom panel) in the spleen of NOD, NOD.NZW-Chr7, and NZW mice. Pooled spleens from B6, NOD, and NZW mice (C) or NOD and NOD.NZW-Chr7 mice (D) were sorted according to the CD11clow Bst-2+B220+ pDC phenotype. Sorted pDC were then stimulated in vitro with CpG ODN 2216. Mean cytokine IFN-α concentration (± SD) from the culture supernatant is shown for one experiment [error bars represent SD for technical replicates, representative of three (C) and two (D) independent experiments]. *p < 0.05, **p < 0.01, ****p < 0.0001.

As the level of pDC was not originally measured in NZW mice, we included these mice in our comparison of NOD and NOD.NZW-Chr7 mice. Unexpectedly, the proportion of pDC in NOD.NZW-Chr7 mice was even lower than in NZW mice (Fig. 6B). This was not the case for the absolute number of pDC, in which both NZW and NOD.NZW-Chr7 mice exhibited a similar number of pDC in the spleen (Fig. 6B). Along with the joint LOD score and segregation analysis described above, these results support the view that genetic interactions between two or more loci contribute to the proportion of pDC because the effect of the NZW-derived congenic interval was increased when placed on the NOD genetic background. These data also corroborate the contribution of distinct modifier loci outside the chromosome 7 interval in the regulation of each trait, namely the proportion and absolute number of pDC.

In addition to variation in the proportion of pDC in different mice, pDC from B6 mice are known to produce significantly less IFN-α than pDC from 129Sv mice, in which the latter exhibits a higher proportion of pDC (4). Similarly, we found that pDC from B6 mice produce less IFN-α than pDC from NOD mice on a per-cell basis (Fig. 6C). To assess whether Pdcc1 affects the production of IFN-α, we compared IFN-α production of pDC sorted from both NOD mice and NOD.NZW-Chr7 congenic mice. pDC from NOD.NZW-Chr7 congenic mice produced similar levels of IFN-α than those from NOD mice on a per-cell basis (Fig. 6D). Similarly, pDC from the NZW parental strain also produced similar levels of IFN-α than those from NOD mice on a per-cell basis (Fig. 6C). These results suggest that Pdcc1 affects the size of the pDC compartment and is unlikely to affect IFN-α production by pDC.

The Pdcc1 locus on chromosome 7, currently defined by linkage and congenic analyses, is relatively large (>90 Mb) and presents with >1000 genes. To restrict the candidate gene search for Pdcc1, we performed bone marrow chimera experiments to determine whether pDC proportion is regulated by cell-intrinsic or -extrinsic hematopoietic factors. The two inbred strains used are B10.Br and NOD.H2k, in which both strains carry the same MHC locus, allowing for the efficient generation of mixed chimerism (25, 26). The B10.Br inbred strain is highly similar to the B6 strain, and both strains present a similar frequency of pDC (compare Fig. 7A with 1B). The NOD and the NOD.H2k congenic strains also exhibited a comparable proportion of pDC (compare Fig. 7A with 1B). This was expected, because the MHC locus on chromosome 17 was not linked to the size of the pDC compartment (Figs. 3, 5). Bone marrow from B10.Br and NOD.H2k mice, which carry the same MHC locus yet a different proportion of pDC, were mixed at a 1:1 ratio in a lethally irradiated F1 (B10.Br × NOD.H2k) recipient and left to reconstitute for 8 wk before analysis. By taking advantage of the CD45 congenic markers for each strain, we found that the B10.Br (CD45.2) and NOD.H2k (CD45.1) bone marrow reconstituted the B cell compartment in relatively equal proportions (Fig. 7B). However, both the proportion and absolute number of pDC originating from the NOD.H2k strain were significantly increased in comparison with those from the B10.Br strain (Fig. 7B). These results demonstrate that the size of the pDC compartment is regulated by cell-intrinsic hematopoietic factors, suggesting that potential candidate genes for Pdcc1 should be expressed in hematopoietic cells.

FIGURE 7.
FIGURE 7.

Analysis of cellular processes that may influence the size of the pDC compartment. (A) Representative plots demonstrating pDC proportion in the spleen of B10.Br and NOD.H2k mice. Percentages are shown. n = 4. (B) B10.Br and NOD.H2k bone marrow were mixed in a 1:1 ratio in F1 (B10.Br × NOD.H2k) lethally irradiated recipient mice and left to reconstitute for 8 wk. The percentage (top panels) and absolute number (bottom panels) of B10.Br (CD45.2+) and NOD.H2k (CD45.1+) cells are shown for B cells (left panels) and pDC (right panels). Data for (A) and (B) were acquired on a BD FACS Calibur flow cytometer. (C) Representative plots showing the degree of BrdU incorporation in Bst-2+ pDC from both B10.Br (top panel) and NOD.H2k mice (bottom panel). (D) The histograms present the expression levels of Bcl-2 (top panel) and active caspase-3 (bottom panel) in CD11clow Bst-2+ cells. Data for (C) and (D) were acquired on an FACS Canto I (BD Biosciences) and are representative of four independent experiments. *p < 0.05. FMO, Fluorescence minus one.

Identifying the specific cellular processes that control the size of the pDC compartment may help further restrict the list of candidate genes. The size of the pDC compartment may be influenced by the rate of pDC proliferation in the different mouse strains. We compared the proliferative rate of pDC in B10.Br and NOD.H2k mice by BrdU labeling. As demonstrated by others (8–10), we find that pDC are relatively quiescent. In addition, we show that BrdU incorporation in pDC was similar in both strains (Fig. 7C), suggesting that the higher number of pDC in the spleen of NOD.H2k mice is not due to an apparent increase in cell cycling. An increase in pDC survival provides an alternative explanation for the increased size in pDC compartment in NOD.H2k mice, relative to B10.Br. However, we find that both Bcl-2 and active caspase-3 levels are present at comparable levels in pDC from both strains (Fig. 7D). Together, these findings suggest that the different size of the pDC compartment between B10.Br and NOD.H2k mice is not due to significant variations in pDC proliferation or apoptotic markers and most likely results from other complex cellular processes.

Discussion

pDC have been shown to contribute to both innate and adaptive immunity, as well as rapidly produce vast amounts of type I IFNs in response to viral Ags (5, 27). Consequently, a more comprehensive understanding of the genetic factors that define the size of the pDC compartment may have relevant therapeutic ramifications. Through an unbiased linkage analysis, we demonstrate that the genetic locus Pdcc1, which is located in the proximal region of chromosome 7, affects the size of the pDC compartment, as both pDC proportion and absolute number are linked to this same locus. Importantly, using NOD.NZW-Chr7 congenic mice, we validated the impact of the Pdcc1 locus in determining the size of the pDC compartment. The distribution of pDC proportion and absolute number in F2.Rag mice, as well as the identification of Pdcc2 and Pdcc3 as modifiers of Pdcc1, also highlight that the size of the pDC compartment is a multigenic trait. Finally, we demonstrate that factors of hematopoietic origin contribute toward defining the size of the pDC compartment.

In this study, we performed a linkage analysis of the proportion and absolute number of pDC in a cohort of F2.Rag mice resulting from a cross between B6.Rag and NOD.Rag mice. The use of Rag-deficient mice provided a means to increase the relative proportion of rare immune cell types, allowing better sensitivity in the linkage analysis. Although NOD mice are prone to spontaneously developing autoimmune diseases, NOD.Rag mice lack both T and B cells necessary for autoimmunity. The NOD.Rag mice thus permitted the analysis of pDC in the absence of underlying inflammation caused by the progression of an autoimmune response. Of interest, the relative difference in the proportion and absolute number of pDC between the B6 and NOD genetic backgrounds is maintained in both Rag-sufficient and -deficient mice, in which the B6 genetic background always presents with fewer pDC relative to NOD regardless of the presence or absence of B and T cells. These results suggest that the relative size of the pDC compartment in various genetic backgrounds is defined, at least in part, independently of the presence of both T and B cells. We further show that the linkage of Pdcc1 to the pDC proportion and absolute number is independent of Rag deficiency. Indeed, NOD.NZW-Chr7 Rag-sufficient mice show a low proportion and absolute number of pDC relative to NOD mice, demonstrating that the proximal region of chromosome 7 defines the size of the pDC compartment even in the presence of T and B cells.

Although dominated by the Pdcc1 locus on mouse chromosome 7, three observations from this study suggest that the size of the pDC compartment is a multigenic trait. Firstly, the F2.Rag cohort represents a normal distribution with regards to both the proportion and absolute number of pDC, indicating that more than one locus contributes to these pDC traits. Secondly, chromosomes 8 and 13 exhibited suggestive linkage to the proportion of pDC and chromosomes 1 and 16 to the absolute number of pDC. Although potential false positives, these suggestive linkages may also be concealed to some degree by Pdcc1, which acts as a dominant locus (24) and will need to be validated using congenic strains. Thirdly, a genetic interaction between Pdcc1 and Pdcc2 was significantly linked to the proportion of pDC, whereas that of Pdcc1 and Pdcc3 was linked to the absolute number of pDC in the spleen. The complex distribution of the pDC compartment size resulting from the interactions among the compound genotypes of Pdcc1, Pdcc2, and Pdcc3 further highlights the dominant effect of Pdcc1 in this multigenic trait.

Candidate genes for Pdcc1, Pdcc2, and Pdcc3 remain to be defined. The bone marrow chimeras demonstrate that the size of the pDC compartment is regulated by cell-intrinsic hematopoietic factors. In addition, we find that neither proliferation of pDC nor the expression level of apoptotic markers in pDC vary between B10.Br and NOD.H2k mice; these parameters are thus unlikely to contribute toward defining the size of the pDC compartment among these strains of mice. Rather, these data suggest that the underlying cellular process defining the size of the pDC compartment may either result from more subtle variations in pDC proliferation or apoptosis not detectable in the context of the assays performed in this study or from more complex hematopoietic-dependent factors, including but not limited to the presence of a key cytokine, cell–cell interaction with other hematopoietic cells, and/or the rate of hematopoietic differentiation of pDC. To that effect, a number of genes (Spib, Tcf4, Irf4, Irf8, and Flt3) have been shown to contribute to the hematopoietic differentiation of pDC (7, 11, 12, 14, 15). Of these genes, Spib and Flt3l (Flt3 ligand) are encoded on chromosome 7 between 51 and 53 Mb, comprised within the NOD.NZW-Chr7 congenic interval contributing to the size of the pDC compartment. Spi-B is a transcription factor implicated in the differentiation of both B cells and pDC (11, 28, 29), in which overexpression of Spi-B in hematopoietic progenitors leads to an increase in pDC number (11). In addition, Flt3L promotes the differentiation of all DC subsets and is known to increase pDC number (30). Spib and Flt3l are thus two candidate genes of interest, which may define the size of the pDC compartment. Additional studies are required to identify which are the causal genetic variants within Pdcc1, Pdcc2, and Pdcc3 responsible for these locus effects.

As we and others have observed variation for IFN-α production from pDC of various mouse strains, we examined the influence of the Pdcc1 locus on IFN-α production by pDC. As for NZW mice, pDC from both NOD and NOD.NZW-Chr7 mice produced similar levels of IFN-α upon in vitro stimulation. These results suggest that the Pdcc1 locus is linked to the size of the pDC compartment at steady state and is unlikely to participate in modulating the production of IFN-α by pDC. Of interest, others have recently shown that serum IFN-α levels determine the size of the pDC compartment in inflammatory conditions (31). It is thus tempting to suggest that different cellular mechanisms control the size of the pDC compartment in steady state and under inflammatory conditions.

Other than their role in the production of IFN-α, pDC have been associated with the induction of immune tolerance, in which pDC favor the differentiation and function of regulatory T cells (32–34). In agreement with a role for pDC in immune tolerance, T1D patients exhibited a lower proportion of pDC in the blood compared with unaffected individuals (35). Notwithstanding, the contribution of pDC in the establishment of self-tolerance and in the prevention of autoimmune disease progression remains controversial (36). Indeed, pDC are found at higher proportion in various autoimmune pathologies, including systemic lupus erythematosus, psoriasis, and T1D (37–39). Moreover, type I IFN responses, which are predominantly due to activated pDCs, are associated with increased susceptibility to systemic lupus erythematosus, Sjogren’s syndrome, and T1D (40–43). In NOD mice, elimination of pDC has been reported to both facilitate the progression to insulitis (44) and prevent diabetes onset (45). Lastly, our study identified a higher proportion of pDC in NOD mice, which spontaneously develop sialadenitis, thyroiditis, and autoimmune diabetes (46). Hence, the contribution of pDC in immune tolerance and prevention of autoimmunity remains to be clearly defined (36).

The data presented in this study suggest that an elevated proportion of pDC is neither necessary nor sufficient for autoimmune diabetes progression. First, NOD.NZW-Chr7 mice present a low proportion of pDC relative to NOD mice, but both strains show a comparable incidence of autoimmune diabetes (17), demonstrating that elevated proportion of pDC is not necessary for autoimmune diabetes progression. Second, both NOR mice, which are highly genetically related to NOD mice (47–49), and 129Sv mice (4) exhibit a high proportion of pDC similar to NOD mice, yet do not develop insulitis or diabetes. This observation indicates that elevated pDC levels are not sufficient to promote T1D pathogenesis. This, however, does not preclude a potential contribution of increased pDC levels in other autoimmune diseases. Indeed, sialadenitis is less severe in NOD.NZW-Chr7 mice (17), which correlates with low pDC proportion. Further investigation of the pathophysiological processes leading to the distinct autoimmune pathologies of insulitis and sialadenitis in NOD mice may better characterize the contribution of pDC to specific autoimmune diseases.

Finally, as pDC have been mostly studied for the ability to effectively produce IFN-α upon viral infections, it would be interesting to investigate the efficiency of NOD and NOR inbred mice, relative to mice presenting a lower proportion of pDC, in the clearance of viral pathogens. Accordingly, elimination of pDC influences the antiviral immune response (5). Therefore, characterization of the Pdcc1, Pdcc2, and Pdcc3 loci could assist in the identification of the molecular determinants that influence the size of the pDC compartment needed to improve antiviral responses and/or immune tolerance regimens.

Disclosures

The authors have no financial conflicts of interest.

Acknowledgments

We thank Sean Wiltshire for support and advice with regards to the use of R software, Lisa Bellemare and Marycruz Domínguez-Punaro for technical support with regards to lung and liver samples, Stacey Fynch and Lorraine Elkerbout for technical support with regards to tissue perfusions, Marie-Josée Guyon and the animal house staff for curating the mouse colonies, Dr. Nathalie Labrecque for critical review of the manuscript, and Erin E. Hillhouse for help with editing the manuscript.

Footnotes

  • This work was supported by grants to S.L. from Diabète Québec, the Canadian Foundation for Innovation, and the Natural Sciences and Engineering Research Council of Canada, as well as grants to T.C.B. from the Juvenile Diabetes Research Foundation and the Victorian Government’s Operational Infrastructure Support Program. A.-N.P. holds scholarships from Diabète Québec and the University of Montreal. F.G.-D. holds a studentship from the Canadian Institutes of Health Research. M.P.A. holds an Australian Postgraduate Award. S.L. holds a New Investigator award from the Canadian Institutes of Health Research.

  • The online version of this article contains supplemental material.

  • Abbreviations used in this article:

    B6
    C57BL/6
    DC
    dendritic cell
    IRF
    IFN regulatory factor
    LOD
    logarithm of odds
    pDC
    plasmacytoid dendritic cell
    Pdcc
    plasmacytoid dendritic cell compartment
    SNP
    single nucleotide polymorphism
    T1D
    type 1 diabetes.

  • Received July 22, 2011.
  • Accepted March 13, 2012.

References

  1. ↵
    1. Colonna M.,
    2. G. Trinchieri,
    3. Y. J. Liu
    . 2004. Plasmacytoid dendritic cells in immunity. Nat. Immunol. 5: 1219–1226.
    OpenUrlCrossRefPubMed
    1. Cao W.,
    2. Y. J. Liu
    . 2007. Innate immune functions of plasmacytoid dendritic cells. Curr. Opin. Immunol. 19: 24–30.
    OpenUrlCrossRefPubMed
  3. ↵
    1. Reizis B.,
    2. A. Bunin,
    3. H. S. Ghosh,
    4. K. L. Lewis,
    5. V. Sisirak
    . 2011. Plasmacytoid dendritic cells: recent progress and open questions. Annu. Rev. Immunol. 29: 163–183.
    OpenUrlCrossRefPubMed
  4. ↵
    1. Asselin-Paturel C.,
    2. G. Brizard,
    3. J. J. Pin,
    4. F. Brière,
    5. G. Trinchieri
    . 2003. Mouse strain differences in plasmacytoid dendritic cell frequency and function revealed by a novel monoclonal antibody. J. Immunol. 171: 6466–6477.
    OpenUrlAbstract/FREE Full Text
  5. ↵
    1. Swiecki M.,
    2. S. Gilfillan,
    3. W. Vermi,
    4. Y. Wang,
    5. M. Colonna
    . 2010. Plasmacytoid dendritic cell ablation impacts early interferon responses and antiviral NK and CD8(+) T cell accrual. Immunity 33: 955–966.
    OpenUrlCrossRefPubMed
  6. ↵
    1. Naik S. H.,
    2. P. Sathe,
    3. H. Y. Park,
    4. D. Metcalf,
    5. A. I. Proietto,
    6. A. Dakic,
    7. S. Carotta,
    8. M. O’Keeffe,
    9. M. Bahlo,
    10. A. Papenfuss,
    11. et al
    . 2007. Development of plasmacytoid and conventional dendritic cell subtypes from single precursor cells derived in vitro and in vivo. Nat. Immunol. 8: 1217–1226.
    OpenUrlCrossRefPubMed
  7. ↵
    1. Wu L.,
    2. Y. J. Liu
    . 2007. Development of dendritic-cell lineages. Immunity 26: 741–750.
    OpenUrlCrossRefPubMed
  8. ↵
    1. Kamath A. T.,
    2. J. Pooley,
    3. M. A. O’Keeffe,
    4. D. Vremec,
    5. Y. Zhan,
    6. A. M. Lew,
    7. A. D’Amico,
    8. L. Wu,
    9. D. F. Tough,
    10. K. Shortman
    . 2000. The development, maturation, and turnover rate of mouse spleen dendritic cell populations. J. Immunol. 165: 6762–6770.
    OpenUrlAbstract/FREE Full Text
  9. ↵
    1. Liu K.,
    2. C. Waskow,
    3. X. Liu,
    4. K. Yao,
    5. J. Hoh,
    6. M. Nussenzweig
    . 2007. Origin of dendritic cells in peripheral lymphoid organs of mice. Nat. Immunol. 8: 578–583.
    OpenUrlCrossRefPubMed
  10. ↵
    1. O’Keeffe M.,
    2. H. Hochrein,
    3. D. Vremec,
    4. I. Caminschi,
    5. J. L. Miller,
    6. E. M. Anders,
    7. L. Wu,
    8. M. H. Lahoud,
    9. S. Henri,
    10. B. Scott,
    11. et al
    . 2002. Mouse plasmacytoid cells: long-lived cells, heterogeneous in surface phenotype and function, that differentiate into CD8(+) dendritic cells only after microbial stimulus. J. Exp. Med. 196: 1307–1319.
    OpenUrlAbstract/FREE Full Text
  11. ↵
    1. Schotte R.,
    2. M. C. Rissoan,
    3. N. Bendriss-Vermare,
    4. J. M. Bridon,
    5. T. Duhen,
    6. K. Weijer,
    7. F. Brière,
    8. H. Spits
    . 2003. The transcription factor Spi-B is expressed in plasmacytoid DC precursors and inhibits T-, B-, and NK-cell development. Blood 101: 1015–1023.
    OpenUrlAbstract/FREE Full Text
  12. ↵
    1. Schotte R.,
    2. M. Nagasawa,
    3. K. Weijer,
    4. H. Spits,
    5. B. Blom
    . 2004. The ETS transcription factor Spi-B is required for human plasmacytoid dendritic cell development. J. Exp. Med. 200: 1503–1509.
    OpenUrlAbstract/FREE Full Text
  13. ↵
    1. Schiavoni G.,
    2. F. Mattei,
    3. P. Sestili,
    4. P. Borghi,
    5. M. Venditti,
    6. H. C. Morse III.,
    7. F. Belardelli,
    8. L. Gabriele
    . 2002. ICSBP is essential for the development of mouse type I interferon-producing cells and for the generation and activation of CD8alpha(+) dendritic cells. J. Exp. Med. 196: 1415–1425.
    OpenUrlAbstract/FREE Full Text
  14. ↵
    1. Cisse B.,
    2. M. L. Caton,
    3. M. Lehner,
    4. T. Maeda,
    5. S. Scheu,
    6. R. Locksley,
    7. D. Holmberg,
    8. C. Zweier,
    9. N. S. den Hollander,
    10. S. G. Kant,
    11. et al
    . 2008. Transcription factor E2-2 is an essential and specific regulator of plasmacytoid dendritic cell development. Cell 135: 37–48.
    OpenUrlCrossRefPubMed
  15. ↵
    1. Ghosh H. S.,
    2. B. Cisse,
    3. A. Bunin,
    4. K. L. Lewis,
    5. B. Reizis
    . 2010. Continuous expression of the transcription factor e2-2 maintains the cell fate of mature plasmacytoid dendritic cells. Immunity 33: 905–916.
    OpenUrlCrossRefPubMed
  16. ↵
    1. Podolin P. L.,
    2. A. Pressey,
    3. N. H. DeLarato,
    4. P. A. Fischer,
    5. L. B. Peterson,
    6. L. S. Wicker
    . 1993. I-E+ nonobese diabetic mice develop insulitis and diabetes. J. Exp. Med. 178: 793–803.
    OpenUrlAbstract/FREE Full Text
  17. ↵
    1. Burt R. A.,
    2. L. Watkins,
    3. I. K. Tan,
    4. N. Wang,
    5. F. Quirk,
    6. L. Mackin,
    7. P. Morgan,
    8. J. G. Zhang,
    9. S. P. Berzins,
    10. G. Morahan,
    11. T. C. Brodnicki
    . 2010. An NZW-derived interval on chromosome 7 moderates sialadenitis, but not insulitis in congenic nonobese diabetic mice. J. Immunol. 184: 859–868.
    OpenUrlAbstract/FREE Full Text
  18. ↵
    1. Vremec D.,
    2. J. Pooley,
    3. H. Hochrein,
    4. L. Wu,
    5. K. Shortman
    . 2000. CD4 and CD8 expression by dendritic cell subtypes in mouse thymus and spleen. J. Immunol. 164: 2978–2986.
    OpenUrlAbstract/FREE Full Text
  19. ↵
    1. Broman K. W.,
    2. H. Wu,
    3. S. Sen,
    4. G. A. Churchill
    . 2003. R/qtl: QTL mapping in experimental crosses. Bioinformatics 19: 889–890.
    OpenUrlAbstract/FREE Full Text
  20. ↵
    1. Herzenberg L. A.,
    2. J. Tung,
    3. W. A. Moore,
    4. L. A. Herzenberg,
    5. D. R. Parks
    . 2006. Interpreting flow cytometry data: a guide for the perplexed. Nat. Immunol. 7: 681–685.
    OpenUrlCrossRefPubMed
  21. ↵
    1. Wicker L. S.,
    2. J. A. Todd,
    3. L. B. Peterson
    . 1995. Genetic control of autoimmune diabetes in the NOD mouse. Annu. Rev. Immunol. 13: 179–200.
    OpenUrlCrossRefPubMed
  22. ↵
    1. Maier L. M.,
    2. L. S. Wicker
    . 2005. Genetic susceptibility to type 1 diabetes. Curr. Opin. Immunol. 17: 601–608.
    OpenUrlCrossRefPubMed
  23. ↵
    1. Blasius A. L.,
    2. E. Giurisato,
    3. M. Cella,
    4. R. D. Schreiber,
    5. A. S. Shaw,
    6. M. Colonna
    . 2006. Bone marrow stromal cell antigen 2 is a specific marker of type I IFN-producing cells in the naive mouse, but a promiscuous cell surface antigen following IFN stimulation. J. Immunol. 177: 3260–3265.
    OpenUrlAbstract/FREE Full Text
  24. ↵
    1. Carlborg O.,
    2. C. S. Haley
    . 2004. Epistasis: too often neglected in complex trait studies? Nat. Rev. Genet. 5: 618–625.
    OpenUrlCrossRefPubMed
  25. ↵
    1. Lesage S.,
    2. S. B. Hartley,
    3. S. Akkaraju,
    4. J. Wilson,
    5. M. Townsend,
    6. C. C. Goodnow
    . 2002. Failure to censor forbidden clones of CD4 T cells in autoimmune diabetes. J. Exp. Med. 196: 1175–1188.
    OpenUrlAbstract/FREE Full Text
  26. ↵
    1. Guimont-Desrochers F.,
    2. Z. J. Cappello,
    3. M. Chagnon,
    4. M. McDuffie,
    5. S. Lesage
    . 2009. Cutting edge: genetic characterization of IFN-producing killer dendritic cells. J. Immunol. 182: 5193–5197.
    OpenUrlAbstract/FREE Full Text
  27. ↵
    1. Asselin-Paturel C.,
    2. A. Boonstra,
    3. M. Dalod,
    4. I. Durand,
    5. N. Yessaad,
    6. C. Dezutter-Dambuyant,
    7. A. Vicari,
    8. A. O’Garra,
    9. C. Biron,
    10. F. Brière,
    11. G. Trinchieri
    . 2001. Mouse type I IFN-producing cells are immature APCs with plasmacytoid morphology. Nat. Immunol. 2: 1144–1150.
    OpenUrlCrossRefPubMed
  28. ↵
    1. DeKoter R. P.,
    2. M. Geadah,
    3. S. Khoosal,
    4. L. S. Xu,
    5. G. Thillainadesan,
    6. J. Torchia,
    7. S. S. Chin,
    8. L. A. Garrett-Sinha
    . 2010. Regulation of follicular B cell differentiation by the related E26 transformation-specific transcription factors PU.1, Spi-B, and Spi-C. J. Immunol. 185: 7374–7384.
    OpenUrlAbstract/FREE Full Text
  29. ↵
    1. Su G. H.,
    2. H. M. Chen,
    3. N. Muthusamy,
    4. L. A. Garrett-Sinha,
    5. D. Baunoch,
    6. D. G. Tenen,
    7. M. C. Simon
    . 1997. Defective B cell receptor-mediated responses in mice lacking the Ets protein, Spi-B. EMBO J. 16: 7118–7129.
    OpenUrlAbstract
  30. ↵
    1. Maraskovsky E.,
    2. K. Brasel,
    3. M. Teepe,
    4. E. R. Roux,
    5. S. D. Lyman,
    6. K. Shortman,
    7. H. J. McKenna
    . 1996. Dramatic increase in the numbers of functionally mature dendritic cells in Flt3 ligand-treated mice: multiple dendritic cell subpopulations identified. J. Exp. Med. 184: 1953–1962.
    OpenUrlAbstract/FREE Full Text
  31. ↵
    1. Swiecki M.,
    2. Y. Wang,
    3. W. Vermi,
    4. S. Gilfillan,
    5. R. D. Schreiber,
    6. M. Colonna
    . 2011. Type I interferon negatively controls plasmacytoid dendritic cell numbers in vivo. J. Exp. Med. 208: 2367–2374.
    OpenUrlAbstract/FREE Full Text
  32. ↵
    1. Huang Y.,
    2. L. D. Bozulic,
    3. T. Miller,
    4. H. Xu,
    5. L. R. Hussain,
    6. S. T. Ildstad
    . 2011. CD8&alpha;+ plasmacytoid precursor DCs induce antigen-specific regulatory T cells that enhance HSC engraftment in vivo. Blood 117: 2494–2505.
    OpenUrlAbstract/FREE Full Text
    1. Ochando J. C.,
    2. C. Homma,
    3. Y. Yang,
    4. A. Hidalgo,
    5. A. Garin,
    6. F. Tacke,
    7. V. Angeli,
    8. Y. Li,
    9. P. Boros,
    10. Y. Ding,
    11. et al
    . 2006. Alloantigen-presenting plasmacytoid dendritic cells mediate tolerance to vascularized grafts. Nat. Immunol. 7: 652–662.
    OpenUrlCrossRefPubMed
  34. ↵
    1. Ito T.,
    2. M. Yang,
    3. Y. H. Wang,
    4. R. Lande,
    5. J. Gregorio,
    6. O. A. Perng,
    7. X. F. Qin,
    8. Y. J. Liu,
    9. M. Gilliet
    . 2007. Plasmacytoid dendritic cells prime IL-10-producing T regulatory cells by inducible costimulator ligand. J. Exp. Med. 204: 105–115.
    OpenUrlAbstract/FREE Full Text
  35. ↵
    1. Chen X.,
    2. L. H. Makala,
    3. Y. Jin,
    4. D. Hopkins,
    5. A. Muir,
    6. N. Garge,
    7. R. H. Podolsky,
    8. J. X. She
    . 2008. Type 1 diabetes patients have significantly lower frequency of plasmacytoid dendritic cells in the peripheral blood. Clin. Immunol. 129: 413–418.
    OpenUrlCrossRefPubMed
  36. ↵
    1. Tisch R.,
    2. B. Wang
    . 2009. Role of plasmacytoid dendritic cells in type 1 diabetes: friend or foe? Diabetes 58: 12–13.
    OpenUrlFREE Full Text
  37. ↵
    1. Bowcock A. M.,
    2. J. G. Krueger
    . 2005. Getting under the skin: the immunogenetics of psoriasis. Nat. Rev. Immunol. 5: 699–711.
    OpenUrlCrossRefPubMed
    1. Farkas L.,
    2. K. Beiske,
    3. F. Lund-Johansen,
    4. P. Brandtzaeg,
    5. F. L. Jahnsen
    . 2001. Plasmacytoid dendritic cells (natural interferon- alpha/beta-producing cells) accumulate in cutaneous lupus erythematosus lesions. Am. J. Pathol. 159: 237–243.
    OpenUrlCrossRefPubMed
  39. ↵
    1. Allen J. S.,
    2. K. Pang,
    3. A. Skowera,
    4. R. Ellis,
    5. C. Rackham,
    6. B. Lozanoska-Ochser,
    7. T. Tree,
    8. R. D. Leslie,
    9. J. M. Tremble,
    10. C. M. Dayan,
    11. M. Peakman
    . 2009. Plasmacytoid dendritic cells are proportionally expanded at diagnosis of type 1 diabetes and enhance islet autoantigen presentation to T-cells through immune complex capture. Diabetes 58: 138–145.
    OpenUrlAbstract/FREE Full Text
  40. ↵
    1. Palucka A. K.,
    2. J. P. Blanck,
    3. L. Bennett,
    4. V. Pascual,
    5. J. Banchereau
    . 2005. Cross-regulation of TNF and IFN-alpha in autoimmune diseases. Proc. Natl. Acad. Sci. USA 102: 3372–3377.
    OpenUrlAbstract/FREE Full Text
    1. Li Q.,
    2. B. Xu,
    3. S. A. Michie,
    4. K. H. Rubins,
    5. R. D. Schreriber,
    6. H. O. McDevitt
    . 2008. Interferon-alpha initiates type 1 diabetes in nonobese diabetic mice. Proc. Natl. Acad. Sci. USA 105: 12439–12444.
    OpenUrlAbstract/FREE Full Text
    1. Nejentsev S.,
    2. N. Walker,
    3. D. Riches,
    4. M. Egholm,
    5. J. A. Todd
    . 2009. Rare variants of IFIH1, a gene implicated in antiviral responses, protect against type 1 diabetes. Science 324: 387–389.
    OpenUrlAbstract/FREE Full Text
  43. ↵
    1. Båve U.,
    2. G. Nordmark,
    3. T. Lövgren,
    4. J. Rönnelid,
    5. S. Cajander,
    6. M. L. Eloranta,
    7. G. V. Alm,
    8. L. Rönnblom
    . 2005. Activation of the type I interferon system in primary Sjögren’s syndrome: a possible etiopathogenic mechanism. Arthritis Rheum. 52: 1185–1195.
    OpenUrlCrossRefPubMed
  44. ↵
    1. Saxena V.,
    2. J. K. Ondr,
    3. A. F. Magnusen,
    4. D. H. Munn,
    5. J. D. Katz
    . 2007. The countervailing actions of myeloid and plasmacytoid dendritic cells control autoimmune diabetes in the nonobese diabetic mouse. J. Immunol. 179: 5041–5053.
    OpenUrlAbstract/FREE Full Text
  45. ↵
    1. Li Q.,
    2. H. O. McDevitt
    . 2011. The role of interferon alpha in initiation of type I diabetes in the NOD mouse. Clin. Immunol. 140: 3–7.
    OpenUrlCrossRefPubMed
  46. ↵
    1. Anderson M. S.,
    2. J. A. Bluestone
    . 2005. The NOD mouse: a model of immune dysregulation. Annu. Rev. Immunol. 23: 447–485.
    OpenUrlCrossRefPubMed
  47. ↵
    1. Serreze D. V.,
    2. M. Prochazka,
    3. P. C. Reifsnyder,
    4. M. M. Bridgett,
    5. E. H. Leiter
    . 1994. Use of recombinant congenic and congenic strains of NOD mice to identify a new insulin-dependent diabetes resistance gene. J. Exp. Med. 180: 1553–1558.
    OpenUrlAbstract/FREE Full Text
    1. Ivakine E. A.,
    2. C. J. Fox,
    3. A. D. Paterson,
    4. S. M. Mortin-Toth,
    5. A. Canty,
    6. D. S. Walton,
    7. K. Aleksa,
    8. S. Ito,
    9. J. S. Danska
    . 2005. Sex-specific effect of insulin-dependent diabetes 4 on regulation of diabetes pathogenesis in the nonobese diabetic mouse. J. Immunol. 174: 7129–7140.
    OpenUrlAbstract/FREE Full Text
  49. ↵
    1. Reifsnyder P. C.,
    2. R. Li,
    3. P. A. Silveira,
    4. G. Churchill,
    5. D. V. Serreze,
    6. E. H. Leiter
    . 2005. Conditioning the genome identifies additional diabetes resistance loci in Type I diabetes resistant NOR/Lt mice. Genes Immun. 6: 528–538.
    OpenUrlCrossRefPubMed
Back to top

In this issue

Print
Download PDF
Article Alerts
Email Article
Citation Tools

Jump to section