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Originally published online as doi:10.1189/jlb.1205739 on July 14, 2006

Published online before print July 14, 2006
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(Journal of Leukocyte Biology. 2006;80:651-658.)
© 2006 by Society for Leukocyte Biology

IP3Rs are sufficient for dendritic cell Ca2+ signaling in the absence of RyR1

Meaghan Stolk*, Matilde Leon-Ponte*, Mia Merrill*, Gerard P. Ahern{dagger},1 and Peta J. O’Connell*,2

* Robarts Research Institute, London, Ontario, Canada; and
{dagger} Georgetown University, Washington, DC

1 Correspondence: Georgetown University, MedDent SW401, 3900 Reservoir Rd., Washington, DC 20007. E-mail: gpa3{at}georgetown.edu


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ABSTRACT
 
Calcium (Ca2+) signaling plays a pivotal role in the function of dendritic cells (DC). The Type 1 ryanodine receptor (RyR), a major intracellular Ca2+ channel, is highly expressed in immature DC. We therefore investigated whether RyR1 plays a role in DC development and function by studying properties of DC derived from wild-type (WT) and RyR1 null [knockout (KO)] mice. Fetal liver cells from WT and RyR1 KO mice retained full hematopoietic competence. Adoptive transfer of these cells into congenic hosts resulted in the generation of functionally equivalent DC populations. WT and RyR1 KO DC exhibited a similar capacity to mature in response to inflammatory and/or activation stimuli, to endocytose antigen, and to stimulate T cell proliferation. Moreover, the absence of RyR1 did not lead to de novo expression of RyR2 or RyR3. WT and RyR KO DC express all three isoforms of inositol 1,4,5-trisphosphate receptor (IP3R), although Type 3 IP3R gene transcripts are predominant. Further, IP3-mediated Ca2+ transients proceed normally after inhibition of RyRs with dantrolene. Signaling via IP3R may therefore be sufficient to drive essential DC Ca2+ signaling processes in the absence of RyR expression or function.

Key Words: calcium • ryanodine receptors • leukocytes • immune function


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INTRODUCTION
 
Calcium (Ca2+) signaling plays a key role in the regulation of immune cell function. A sustained increase in intracellular Ca2+ ([Ca2+]i) accompanies T and B cell receptor signaling and is necessary for interleukin-2 (IL-2) production, cellular proliferation, and antibody secretion [1 , 2 ]. Similarly, many critical processes and functions in professional antigen-presenting cells, such as dendritic cells (DC), appear to involve Ca2+ signaling. DC maturation, including the enhanced expression of major histocompatibility complex (MHC) Class II and costimulatory molecules, is inhibited by chelation of external Ca2+ [3 ]. Conversely, agents that mobilize [Ca2+]i can promote DC maturation in the absence of typical maturation-inducing cytokines or stimuli [3 4 5 ]. An increase in free [Ca2+]i is essential for the uptake and processing of apoptotic bodies [6 ]. Similarly, stimuli that raise [Ca2+]i trigger the secretion of preassembled cytokines and inflammatory molecules from vesicular stores [7 8 9 10 ]. Chemotactic molecules and T cell-derived signals uniformly produce Ca2+ increases in DC [11 12 13 14 15 ], suggesting that Ca2+ transients regulate DC migration and their capacity to initiate adaptive immune responses.

The important role of Ca2+ in all these functions emphasizes the need to better understand the Ca2+ signaling pathways in DC. Many of the G-protein-coupled receptors and receptor protein tyrosine kinases expressed by DC induce Ca2+ mobilization. These signal via phospholipase C (PLC)ß and PLC{gamma}, respectively, leading to formation of inositol 1,4,5-trisphosphate (IP3) and Ca2+ release from IP3 receptors (IP3Rs) in the endoplasmic reticulum [16 ]. Another major class of [Ca2+]i channel found in many cell types is the ryanodine receptors (RyRs). Although their best-known function is in muscle contraction, RyRs may play important roles in diverse Ca2+ signaling pathways [17 18 19 ]. Three RyR types (RyRs1–3) have been described; each is the product of a different gene. RyR1 and RyR2 channels are expressed predominantly in skeletal muscle and heart, respectively, and RyR3 is more widely expressed. RyR3 expression has been documented in human T cells and may contribute to the sustained Ca2+ rise that is necessary for IL-2 activation and T cell proliferation [20 , 21 ]. B lymphocytes have recently been demonstrated to express RyR1 [22 , 23 ]. All RyRs are triggered by small increases in [Ca2+]i (>0.5 µM), which in turn, lead to a much greater [Ca2+]i rise (a process termed Ca2+-induced Ca2+ release). In this way, RyRs can markedly alter the amplitude and duration of [Ca2+]i transients. Indeed, this mechanism may underlie the role of RyRs in leukocytes and other nonexcitable tissues.

We have recently identified the expression of functional RyR1 channels in mouse bone marrow-derived DC (BMDC) [24 ]. RyR ligands were found to induce robust Ca2+ rises in DC, and the expression of RyR1 gene transcripts is regulated developmentally. Immature myeloid DC show marked RyR1 expression (comparable with skeletal muscle), but the expression of RyR1 RNA is reduced dramatically upon maturation [24 ]. We hypothesized that RyRs in these cells may serve to amplify Ca2+ rises initiated by IP3Rs and trigger signaling pathways critical for DC development or function. Here, we have characterized the properties of DC derived from wild-type (WT) and RyR1 knockout (KO) mice. We show that WT and RyR1 KO DC are functionally equivalent in their ability to mature in response to inflammatory and/or activation stimuli and their capacity to endocytose antigen and stimulate T cells. Moreover, the absence of RyR1 did not lead to de novo expression of RyR2 or RyR3. Thus, we conclude that RyR1 alone is not essential for these DC functions. Signaling via IP3Rs, of which Type 3 is expressed predominantly by DC, appears to be sufficient to mediate [Ca2+]i signaling in DC.


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MATERIALS AND METHODS
 
Animals
Male C57BL (C57) or BALB/c mice, 7–12 weeks of age, were purchased from Charles River Laboratory (Wilmington, MA) and housed in the Health Sciences Animal Facility, University of Western Ontario (Canada). CD45.1 congenic mice [(B6.SJL-PtprcaPep3b/BoyJ (CD45.1+)], 6–8 weeks of age, were obtained from The Jackson Laboratory (Bar Harbor, ME). Mice heterozygous for an insertion deletion of RyR1 were obtained from Dr. Paul D. Allen (Brigham and Women’s Hospital, Boston, MA) [25 ]. RyR1 mutant mice were housed under specific pathogen-free conditions in the Health Sciences Animal Facility, University of Western Ontario, and backcrossed to the C57Bl/6 background for at least six generations. Mice were genotyped by polymerase chain reaction (PCR; Fig. 1A and 1B ). Genomic DNA was extracted from tail snips using the AquaPure genomic DNA isolation kit (Bio-Rad Laboratories, Hercules, CA). DNA was amplified using Platinum HiFi Taq (Invitrogen, Carlsbad, CA). A 232-bp product was amplified from the WT allele using RyR1-F (5'-GGG AAC CCT AGA AGA GAG ATG ACT G-3') and RyR1-R (5'-CCT TGG TGT GGG CCT TGC TGG-3') primers. The mutant allele was identified by amplification of a 258-bp product using RyR1-F and KpnI-R (5'-CCT GAA GAA CGA GAT CAG CAG CCT CTG TTC C-3'). DNA was amplified by 35 cycles of denaturation at 95°C for 20 s, annealing at 60°C for 30 s, and extension at 72°C for 30 s.


Figure 1
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  		Figure 1. Interruption of the mouse RyR1 gene. (A) Schematic representation of the RyR1 WT gene compared with the RyR1 mutant gene generated by insertion of a neomycin cassette at the KpnI site. Primer positions for binding to RyR1 WT (RyR1-F+RyR1-R) and RyR1 KO (RyR1-F+KpnI-R) genes are indicated. (B) Genomic PCR from Days 12–14 of gestation (E12–14) mouse tail DNA following amplification with RyR1 WT or RyR1 KO primers. Het, heterozygote.

All mice were provided with rodent chow and water ad libitum and used in accordance with the guidelines set forth by the Canadian Council on Animal Care and the Georgetown University Animal Care and Users Committee (Washington, DC).

Reverse transcriptase (RT)-PCR
RNA was extracted using TRIzol (Invitrogen), and first-strand cDNA synthesis was performed using Advantage RT-for-PCR (BD Biosciences, San Jose, CA). For conventional RT-PCR, cDNA was amplified using Taq DNA polymerase (Eppendorf, Hamburg, Germany). The primers are as follows: RyR1 (forward 5'-CTG GCT GTG AAG AAG GCT TTG TGA CTG GAG-3' and reverse 5'-CGT CAG TGA CAA CGC ATC ATC CAT GTG ACC TT-3'), RyR2 (forward 5'-CAC AGA CAA TTC CTT CCT CTA CCT A-3' and reverse 5'-AAC ACC TCT CTT GGT ACA TCT TCC-3'), RyR3 (forward 5'-AGG TTC CTT GCT CTG TTT GT-3' and reverse 5'-TGC TTT GGC CTC TTC TAC TG-3'), IP3R1 (forward 5'-AAG CGG ATG GAC CTG GTG TTA GAA CTG-3' and reverse 5'-AAT TTG TGC TGT GTG CTT CGC GTA GAA CT-3'), IP3R2 (forward 5'-CTG TTC TTC TTC ATC GTC ATC ATC ATC G-3' and reverse 5'-GAA ACC AGT CCA AAT TCT TCT CCG TGA-3'), IP3R3 (forward 5'-CTT CTT TAT CGT CAT CAT CAT CGT GTT G-3' and reverse 5'-AGG TTC TTG TTC TTG ATC ATC TGA GCC A-3'), glyceraldehyde 3-phosphate dehydrogenase (GAPDH; forward 5'-GCC GCC TGG AGA AAC CTG CCA AGT-3' and reverse 5'-TAT TCA AGA GAG TAG GGA GGG CTC-3'), and 18S (forward 5'-ATA ATG CTG ACG CCG CCC AAC ACC-3' and reverse 5'-TGA GAG ACC CAG TAC CGG CTT TCC TA-3'). Transcripts were amplified by 32–40 cycles of cDNA denaturation (20 s at 95°C), followed by 30 s of primer annealing and 30 s extension at 72°C. Annealing temperatures were as follows: RyR1 (63°C), RyR2 (57°C), RyR3 (57°C), IP3R1 (59°C), IP3R2 (56°C), IP3R3 (56°C), GAPDH (57°C), and 18S (56°–59°C). PCR products were resolved as single bands by agarose gel electrophoresis and visualized with SYBR Safe (Molecular Probes, Eugene, OR) fluorescence.

For quantitative PCR (qPCR), cDNA was amplified using iQTM SYBR® Green Supermix (Bio-Rad Laboratories) in an MJ Research Chromo4 system (Bio-Rad Laboratories). All reactions were performed in triplicate, and cycling conditions were as described earlier for conventional RT-PCR. The expression of IP3R1–3 was normalized to the housekeeping gene 18S using the comparative threshold cycle, as described [26 ]. The expression of IP3R gene transcript by BMDC is presented relative to the positive control (brain).

Fetal liver hematopoietic cells and adoptive transfer
Timed pregnancies of mice heterozygous for the RyR1 insertion deletion were used to obtain embryos at E12–14. Fetal livers were removed carefully and disaggregated. Erythrocytes were lysed by hypotonic shock using 0.15 M NH4Cl. For adoptive transfer, adult CD45.1 congenic hosts were sublethally irradiated (500 rads). The following day, hosts were anaesthetized with ketamine [100 mg/kg, intraperitoneally (i.p.)], and xylazine (5 mg/kg, i.p.) and CD45.2+ WT or RyR1 KO fetal liver cells (3.5x106) were transferred by intravenous injection in the retro-orbital sinus. Peripheral blood was collected at weekly intervals following adoptive transfer to monitor engraftment of CD45.2+ cells by monoclonal antibody (mAb) labeling and flow cytometry.

BMDC culture
BM cells were isolated from the femurs and tibias of C57Bl/6 or C57Bl/10 mice and cultured for 5 days at 3 x 105 cells in RPMI 1640 (Invitrogen) supplemented with 10% v/v fetal calf serum (FCS; Hyclone, Logan, UT), nonessential amino acids, L-glutamine, sodium pyruvate, penicillin-streptomycin, 2-mercaptoethanol (all from Invitrogen; referred to subsequently as complete medium), granulocyte macrophage-colony stimulating factor, and IL-4, as described previously [4 , 27 ].

Antibodies and flow cytometry
For mAb labeling, nonspecific binding was blocked by incubating cells with 5% v/v normal goat serum (20 min, 4°C, Sigma Chemical Co., St. Louis, MO) and then washed with phosphate-buffered saline + 0.1% bovine serum albumin. For analysis of hematopoietic cells, E12–24 fetal liver cells were labeled with lineage-specific fluorescein isothiocyanate (FITC) mAb to detect CD3 (17A2), CD4 (L3T3RM4-5), CD5 (53-6.3), CD8 (53-6.7), CD45R (RA3-6B2), TER119, and Gr-1 (RB6-8C5), phycoerythrin (PE) anti-CD90.2 (30-H12), PE-Cy5.5 anti-CD117 (c-kit; 2B8), and biotinylated anti-stem cell antigen (Sca)-1 (D7), detected by secondary labeling with streptavidin-allophycocyanin (SA-APC). BMDC were labeled with anti-CD45.2 FITC (104), biotinylated anti-CD45.1 (A20), followed by SA-peridinin chlorophyll protein (PerCP) Cy5.5, anti-CD11c APC (HL3), and PE-conjugated anti-CD86 PE (GL1) or anti-MHC Class I PE (28-14-8). For analysis of spleen DC subpopulations single-cell suspensions were prepared and labeled with anti-CD45.2 FITC, anti-CD8 PE, and anti-CD11c APC or anti-CD45.2 FITC, anti-CD11c PE, and anti-CD45RB APC. Host T and B cells were excluded by labeling with PerCP Cy5.5-conjugated anti-CD19 (MB19-1) together with biotinylated anti-CD3 and anti-CD45.1, which were detected by SA-PerCP Cy5.5. Antibodies were purchased from Biolegend (San Diego, CA) or BD Biosciences. Labeled cells were fixed with 1% w/v paraformaldehyde and analyzed using a FACSCalibur flow cytometer with Cell Quest software (BD Biosciences). Isotype-matched, irrelevant immunoglobulin (Ig) were used as negative controls.

Endocytosis assay
BMDC were isolated from BM after 4 days, and nonspecific labeling was blocked using 10% v/v FCS and normal goat serum (20 min, 4°C). BMDC were incubated with 0.5 µg/ml dextran-FITC (Sigma Chemical Co., 60 min, 37°C). Incubation in the presence of 10 mM EDTA or at 4°C was used to control for nonspecific labeling. Cells were washed twice and labeled with anti-CD86 PE, biotinylated anti-CD45.1 followed by SA-PerCP Cy5.5 and anti-CD11c APC. Dextran-FITC uptake by RyR1 WT or Ry1 KO was analyzed by flow cytometry.

IL-12 enzyme-linked immunosorbent assay (ELISA) and mixed leukocyte reacation
WT or RyR1 KO BMDC were purified to >95% by cell sorting of CD11c+ CD45.1– cells from Day 5 BM cultures. Sorted WT or RyR1 KO BMDC were cultured for 2 days with lipopolysaccharide (LPS; 50 ng/ml, L2654, Sigma Chemical Co.) or cross-linked anti-CD40 mAb (5 µg/ml, HM40-3, BD Biosciences). Culture supernatants were analyzed by ELISA for IL-6 and IL-12 (Biolegend).

To assess their stimulatory capacity, graded numbers of sorted WT or RyR1 KO BMDC were placed in the wells of a 96-well, round-bottom plate and cocultured with CD3+ T cells (2x105 cell/well), negatively enriched from BALB/c spleens (SpinSepTM T cell enrichment kit, Stem Cell Technologies, Vancouver, Canada). Bulk spleen cells (2x105), allogeneic (C57Bl/6) or syngeneic to the BALB/c T cells, were used as control stimulators. Cultures were incubated for 72 h in a humidified atmosphere of 5% CO2 in air. [3H] Thymidine (TdR; 1 µCi) was added to each well for the final 18 h of culture. Cells were harvested using a multiple well harvester, and [3H] TdR incorporation was determined by scintillation counting. Results are expressed as the mean counts per minute (cpm) ±1 SD from triplicate cultures.

Ca2+ imaging
BMDC and sensory neurons were loaded with 1 µM Fluo 4-AM (Molecular Probes) for 20 min and washed for a further 10–20 min prior to recording. The dye was excited at 480 ± 15 nm. Emitted fluorescence was filtered with a 535 ± 20-nm bandpass filter, captured by a SPOT RT digital camera (Diagnostic Instruments, Sterling Heights, MI), and read into a computer. Analysis was performed offline using Simple PCI software (Compix Inc., Sewickley, PA). Adenosine 5'-triphosphate (ATP; 100 µM) was applied via a gravity-fed micropipette (~100 µm diameter) positioned at a distance of ~0.5 mm from the cell of interest. Substance P (SP; 50 ng/ml) was applied by a puffer pipette (1–2 µm diameter) located 50 µm from individual cells, which were pretreated with the PLC inhibitor U73122 (10 µM, Calbiochem, San Diego, CA) or the RyR inhibitor dantrolene (10 µM, Sigma Chemical Co.) for 10 min prior to recording.


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RESULTS
 
RyR1 KO mice are hematopoietically sufficient
To explore the role of RyR1-mediated Ca2+ signaling in the development of hematopoietic progenitors and DC differentiation, we isolated liver cells from fetal mice at E12–14. Expansion of the mouse hematopoietic population occurs primarily in the fetal liver between Days 11 and 15 of gestation. These fetal liver hematopoietic cells then seed the BM and spleen with multipotent progenitors, which later differentiate into mature leukocytes, including DC. Using RT-PCR, RyR1 was identified as the predominant RyR gene expressed by fetal liver hematopoietic cells (Fig. 2A ). It is important that in the genetic absence of RyR1, up-regulation of RyR2 or RyR3 was not observed (Fig. 2A) . To assess the impact of RyR1 on hematopoietic development, we analyzed E12–14 fetal liver cells for a phenotype consistent with that described previously for mouse hematopoietic stems cells [28 ]. Lineage–, CD90lo (Thy-1), CD117+ (c-kit), and Sca-1+ hematopoietic stem cells were identified in RyR1 WT and RyR1 KO fetal liver cells (Fig. 2B) . Thus, deletion of RyR1 does not negatively affect the development of hematopoietic stem cells.


Figure 2
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  		Figure 2. Deletion of RyR1 does not impair the development of cells with the phenotype of hematopoietic stem cells. (A) Expression of RyR subtypes by E12–14 fetal liver hematopoietic cells was determined by RT-PCR of sequences specific for each of the three RyRs (RyR1–3). Skeletal muscle, heart, and brain samples were used as positive controls for RyR1–3 expression, respectively. Comparable quantities of cDNA were ensured by amplification of GAPDH. (B, Top) E12–14 fetal liver cells from WT or RyR1 KO mice were gated as lineage-negative (CD3, CD4, CD5, CD8, CD45RB, Ter 119, and Gr-1) and CD90.2 (Thy-1.2)lo. (Middle) Gated cells were then analyzed for their expression of CD117 (c-kit) and Sca-1. (Bottom) Isotype-matched Ig was used as negative controls.

DC differentiation and maturation are not impaired by the absence of RyR1
We have previously shown that RyR1 is expressed by immature mouse BMDC and is down-regulated on maturation; thus, we next asked whether RyR1 is necessary for DC differentiation. Homozygous deletion of RyR1 is a birth lethal genotype. Thus, to examine the development and function of RyR1-deficient DC, E12–14 fetal liver hematopoietic cells were isolated from WT and RyR1 KO mice and reconstituted into sublethally irradiated CD45.1 congenic hosts. Reconstitution of the hematopoietic compartment was determined by monitoring the phenotype and frequency of CD45.1+ (host) and CD45.2+ (RyR1 WT or RyR1 KO) peripheral blood leukocytes (data not shown). Spleens were isolated following reconstitution, and their DC constituents were examined by mAb labeling and flow cytometry. Splenic DC were identified by their expression of the mouse DC marker CD11c. DC, which developed from WT or RyR1 KO fetal liver cells, were identified by their expression of CD45.2, and host DC were excluded by the expression of congenic CD45.1. The frequency of splenic CD45.2+ CD11c+ DC was relatively equal and within the expected range of ≤0.5% total leukocytes (range 0.12–0.23%) in mice reconstituted with WT or RyR1 KO fetal liver cells. Moreover, the distribution of mouse interstitial DC subtypes was also similar. As shown in Figure 3A , CD8 DC (CD8{alpha}+ CD11c+), CD11b DC (CD8{alpha}– CD11c+), and plasmacytoid DC (B220+ CD8{alpha}+ CD11c+) were equivalent in the presence and absence of RyR1.


Figure 3
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  		Figure 3. Absence of RyR1 does not impair the development of DC. CD45.1+ C57Bl/6 mice were reconstituted with congenic E12–14 fetal liver cells from CD45.2+ WT or RyR1 KO mice. (A) Spleen cells were gated as CD45.2+-FITC and excluded all cells labeled with biotinylated anti-CD3 and anti-CD45.1 (detected by SA-PerCP Cy5.5). Dot plots show the frequency of CD8 DC (CD8{alpha}+ CD11c+), CD11b DC (CD8{alpha}– CD11c+), and B220 DC (B220+ CD11c+) subtypes from WT versus RyR1 KO mice. Data are representative of three independent experiments. (B) DC were cultured from BM for 4 days. BM cultures were then labeled with mAb and gated as CD45.2+-FITC, excluding cells labeled with biotinylated CD45.1 (detected by SA-PerCP Cy5.5). Dot plots show the expression of CD86 or MHC Class II (IAb) PE by CD45.2+ DC labeled with anti-CD11c APC. Isotype-matched, irrelevant Ig was used to control for nonspecific labeling. (C) RT-PCR of sequences specific for each of the three RyRs (RyR1–3) demonstrates that RyR1 is the subtype expressed by BMDC and is absent in RyR1 KO mice. Skeletal muscle, heart, and brain samples were used as positive controls for RyR1–3 expression, respectively. Data are one of two similar experiments.

Next, we examined the in vitro differentiation of RyR1-deficient DC using a well-characterized protocol for their culture from mouse BM [27 ]. Similar to splenic DC development, the in vitro differentiation of mouse BMDC from RyR1 KO BM was not impaired, compared with the WT control (Fig. 3B) . The frequency of CD11c+ BMDC was similar in RyR1 WT and RyR1 KO BM cultures (range 18.4–19.8%). Moreover, the distribution of immature (MHC Class IIlo, CD86lo CD11c+) and mature BMDC (MHC Class IIhi, CD86hi CD11c+) in BM cultures was also comparable (Fig. 3B) . Further, using RT-PCR, we confirmed our earlier finding that RyR1 is the predominant subtype expressed by BMDC [24 ]. Significantly, in the absence of RyR1, up-regulation of RyR2 or RyR3 was not observed in BMDC (Fig. 3C) . Thus, the absence of Ca2+ signaling via RyR1 does not impede the differentiation of endogenous spleen DC subtypes or in vitro-generated BMDC.

RyR1-deficient BMDC are functionally competent
Having demonstrated that DC differentiation and phenotypic maturation are not impaired by the deletion of RyR1, we next examined if RyR1 KO DC were functionally competent. As professional antigen-presenting cells, DC serve to capture and transport antigen to draining lymph nodes for presentation to T cells. Thus, we first assessed the capacity of WT or RyR1 KO BMDC to endocytose the model antigen, dextran-FITC. As described earlier, BMDC were cultured from congenic hosts reconstituted with RyR1 WT or KO fetal liver cells. DC of host origin were excluded from analysis by their expression of CD45.1. As shown in Figure 4A , immature WT and RyR1 KO BMDC exhibited efficient and approximately equivalent capacity for the uptake of dextran-FITC. In contrast, mature RyR1 WT and RyR1 KO BMDC were poorly endocytic (data not shown). Labeling was inhibited at 4°C or in the presence of EDTA (10 mM), indicating that dextran was actively endocytosed rather than bound to the cell surface.


Figure 4
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  		Figure 4. Absence of RyR1 does not impair the endocytic or stimulatory capacity of BMDC. DC were cultured from the BM of CD45.1+ C57Bl/6 mice reconstituted with congenic E12–14 fetal liver cells from CD45.2+ WT or RyR1 KO mice. (A) After 4 days of culture, DC were incubated with dextran-FITC (0.5 µg/mL), where its uptake by CD11c+ CD45.1– DC at 37°C (filled histogram) was compared with uptake at 4°C (bold histogram) or in the presence of 10 mM EDTA (dotted histogram). (B and C) CD11c+ CD45.1– BMDC were purified by cell sorting. (B) WT or RyR1 KO BMDC were cultured for 2 days with LPS (50 ng/ml), cross-linked anti-CD40 mAb (5 µg/ml), or left untreated. The concentration of IL-6 and IL-12 in culture supernatants was determined by ELISA. (C) Graded numbers of RyR1 WT or RyR1 KO BMDC were cocultured with allogeneic (H2d) CD3+ T cells (1x106/ml) for 72 h. Bulk spleen cells syngeneic (BALB/c) or allogeneic (C57Bl/6) with purified T cells were used as control stimulators. Data are means ± 1 SD from triplicate assays and are representative of two independent experiments.

Next, we assessed the production of proinflammatory cytokines by BMDC and their capacity to stimulate T cell proliferation. CD45.2+ CD11c+ BMDC were purified by cell sorting and cultured with anti-CD40 mAb to simulate ligation by the T cell molecule CD154 (CD40 ligand) or with LPS. Phenotypic maturation, as measured by increased expression of CD86 and cell surface MHC Class II, was observed in WT and RyR1 KO BMDC (data not shown). Relative to untreated BMDC, LPS induced significant and equivalent increases in the secretion of IL-6 and IL-12 into the culture supernatant (approximately ten- and 20-fold, respectively) by WT and RyR1 KO BMDC (Fig. 4B) . Similar, but less pronounced, effects were observed in response to receptor cross-linking with anti-CD40 mAb (Fig. 4B) . The stimulatory capacity of BMDC was assessed by coculture for 3 days with allogeneic CD3+ T cells. WT and RyR1 KO BMDC induced potent and equivalent proliferation of allogenic T cells in vitro (Fig. 4C) and were approximately 12-fold more efficient compared with bulk spleen cells.

IP3R expression and function in BMDC
A potential explanation for the absence of a phenotype in RyR null DC may be provided by compensatory up-regulation of IP3R signaling. To confirm functional IP3R signaling, we performed Ca2+ imaging in response to stimulation with extracellular ATP (100 µM), which is known to activate P2Y receptors. Under control conditions, ATP evoked large Ca2+ transients in nearly all cells tested (Fig. 5A ). In contrast, these ATP-induced responses were inhibited after pretreatment with the PLC inhibitor U73122 (10 µM; Fig. 5B ). These data are therefore consistent with a PLC-IP3-IP3R-mediated signaling pathway and agree with our previous observations that ATP-evoked Ca2+ transients persist in cells bathed in a Ca2+-free medium [4 ].


Figure 5
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  		Figure 5. Demonstration of functional IP3R-mediated Ca2+ signaling in BMDC. (A) Extracellular ATP (100 µM) evoked marked Ca2+ transients in control cells (n=17), but responses to ATP were largely inhibited in cells pretreated with the PLC inhibitor U73122 (10 µM, 10 min, n=8; B). These data are consistent with activation of P2Y receptors coupled to PLC and subsequent IP3R activation. F/F0, F = fluorescence intensity, and F0 = F at rest. (C and D) WT and KO BMDC were examined for IP3R1, IP3R2, and IP3R3 gene transcripts using PCR for isoform-specific sequences. qPCR are means from triplicate results ± 1 SD. Expression of IP3R gene transcript was normalized (18S) and expressed relative to the positive control (brain).

To identify IP3R expression in DC, we performed RT-PCR for the three known isoforms using whole brain as the positive control. As shown in Figure 5C , BMDC obtained from WT mice express mRNA for all three IP3R subtypes. This is consistent with the expression pattern observed in T cells [29 , 30 ]. Significantly, qPCR reveals that the Type 3 IP3R is the predominant gene transcript expressed by BMDC and is present in relatively equivalent quantities in WT and KO BMDC (Fig. 5D) . These data suggest that changes in the amount or pattern of IP3R expression do not occur in the DC in the absence of RyR1.

Previously, we have postulated that RyR1 may serve to amplify IP3-mediated Ca2+ release in DC [24 ]. Treating BMDC with ryanodine elicits Ca2+ release and reduces subsequent ATP-evoked Ca2+ transients [24 ]. This suggests that RyRs and IP3Rs may share the same Ca2+ stores. Colocalization of these receptors may allow for functional interactions, whereby Ca2+ release through IP3Rs in turn elicits further release from nearby RyRs. To explore this possibility, we measured IP3-mediated Ca2+ transients under control conditions or in the presence of the RyR antagonist dantrolene, known to inhibit RyR signaling in BMDC [24 ]. IP3 signaling in BMDC was stimulated by SP [31 ], and Ca2+ responses were normalized to the maximal signal produced by ionomycin (Fig. 6A ). Figure 6 B and C , shows that the mean peak Ca2+ rise evoked by SP was ~50% of that produced by ionomycin, and this was not altered significantly by dantrolene treatment (P=0.12). Thus, RyRs do not appear to contribute to acute IP3-triggered Ca2+ mobilization in BMDC.


Figure 6
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  		Figure 6. RyRs do not amplify IP3R-mediated Ca2+ transients. (A) A Ca2+ transient in a BMDC evoked by SP (50 ng/ml) and subsequent maximal Ca2+ response generated by ionomycin (Iono; 100 µM). (B) Representative Ca2+ transients produced by SP under control conditions or in the presence of the RyR inhibitor dantrolene (10 µM). (C) Mean peak Ca2+ responses evoked by SP (expressed as a percent of ionomycin) for control (n=18) and dantrolene (n=17) conditions (P=0.12, Student’s t-test).


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DISCUSSION
 
Ca2+ signaling plays an important role in DC development and function. Many of the G-protein-coupled receptors and receptor tyrosine kinases expressed by DC induce Ca2+ mobilization. These receptors are often developmentally regulated. Previously, we reported that expression of the Ca2+ channel RyR1 is tightly coupled to mouse BMDC of an immature maturation state [24 ]. In contrast, monocytes and macrophages, which share some developmental and functional features with DC, do not show evidence of RyR1 expression [32 ]. Taken together, these findings suggest a role for RyR1 in DC function or development. We hypothesized that RyR1 expression during hematopoiesis may be important in selecting a DC differentiation pathway, in the function of immature DC, such as antigen uptake and processing, or in terminal maturation and T cell activation. All of these functions depend on Ca2+ signaling.

RyR1 null mice were used to probe the role of RyR signaling in BMDC. Fetal liver cells (E12–14) from RyR1 KO mice reconstituted the hematopoietic compartment of immune-ablated mice with equal efficiency compared with the WT controls. The frequency and distribution of T and B lymphocytes and myeloid lineage monocytes and granulocytes were restored with similar kinetics (data not shown). These findings are consistent with our observations that hematopoietic stem cells are not deficient in RyR1 null mice (see Fig. 2B ). The development of DC subpopulations in vivo and their differentiation in vitro from BM progenitors also occurred similarly to WT controls. Thus, although the Type 1 RyR is the sole member of this Ca2+ channel class expressed by E12–14 fetal liver cells, its deletion does not impede the differentiation of hematopoietic progenitors in vivo nor their capacity to reconstitute the hematopoietic compartment.

Next, we examined whether the absence of RyR1 leads to impaired function or maturation of BMDC. Central to their role as professional antigen-presenting cells, DC are highly specialized for the uptake and processing of foreign and self-antigen and the activation of naïve T cells. However, our data demonstrate that RyR1 KO BMDC do not exhibit impaired endocytic capacity or T cell stimulatory activity. Moreover, RyR1 KO BMDC undergo similar maturation, as determined by expression of the MHC Class II and CD86 (data not shown) and the production of IL-12 compared with WT BMDC. Thus, our data clearly demonstrate that RyR1 signaling is not required for these fundamental aspects of DC biology, as determined in vitro; however, we cannot exclude the possibility that RyR1 signaling contributes to immune functions only discernable in vivo. We considered the possibility that the normal development and function of hematopoietic progenitors and DC in the absence of RyR1 may result from the compensatory up-regulation of alternate RyR isoforms, RyR2 or RyR3. Indeed, a recent study of RyR expression in leukocytes has demonstrated that RyR2 transcripts may be induced in peripheral blood mononuclear cells in response to various stimuli, including hematopoietic growth factors, chemokines, and mitogens [32 ]. The Type 3 RyR has not been detected in peripheral blood cells or primary T cells [32 ], although it is expressed by the immortalized T cell lines Jurkat and BW 5147 [20 , 33 ]. RT-PCR did not reveal de novo expression of RyR2 or RyR3 in RyR1 KO BMDC.

IP3Rs are fundamental [Ca2+]i channels ubiquitously expressed in mammalian cells [16 ]. Our previous report indicated that IP3Rs and RyRs in DC share the same [Ca2+]i store, thereby indicating a functional interplay between RyRs and IP3Rs during DC Ca2+ signaling [24 ]. Many of the chemokines and growth factors that control DC maturation and migration act via G-protein-coupled receptors and initiate IP3-mediated Ca2+ rises. We therefore hypothesized that RyR1 coupled to IP3R Ca2+ stores may serve to amplify IP3-mediated Ca2+ transients. However, in this study, we found that inhibition of RyRs did not reduce Ca2+ transients evoked by SP [31 ]. Three types of IP3Rs have been identified, each encoded from a separate gene. Although numerous studies have described presumptive IP3R signaling in DC, such as Ca2+ transients evoked by chemokines and receptor tyrosine kinase activation, little formal proof of IP3R signaling has been presented. Moreover, IP3R expression and the identity of IP3R isoforms have not been reported for DC. Our data show that BMDC express all three IP3R isoforms; however, IP3R3 is the predominant subtype. The significance of this expression pattern is unclear. Although initial studies suggested that IP3R subtypes differed in their Ca2+ regulation, subsequent studies have found no clear difference [16 ]. However, there are clear tissue-specific differences in IP3R subtype expression. For example, the cerebellum exclusively expresses the type 1 IP3R; accordingly, mice lacking this receptor exhibit severe neurological deficits [34 ]. In addition, the pancreas and salivary glands appear to express Types 2 and 3 receptors exclusively, and mice lacking both these genes have impaired exocrine function [35 ]. In contrast, IP3Rs appear to be ubiquitously expressed in cells of the immune system. For example T cells and B cells express all three subtypes of IP3R [29 , 30 , 36 ]. Although it was originally reported that the Type 1 receptor was essential for T cell receptor-mediated signaling in Jurkat T cells [37 ], a more recent study reported no such defect in thymocytes and splenic T cells from IP3R1 null mice [30 ]. It is important that T cells from IP3R1 null mice are able to mobilize [Ca2+]i and exhibit normal proliferative responses in the absence of compensatory up-regulation of Types 2 and 3 receptors. These data are strikingly similar to the current observations regarding DC and combined, suggest a substantial degree of functional redundancy for [Ca2+]i mobilization in lymphocytes and BMDC.

Our data suggest that efficient signaling through multiple IP3R subtypes in DC may minimize the requirement for RyR signaling. Indeed, we have previously estimated that IP3R-mediated signaling in BMDC stimulated by ATP produces a large increase in the global [Ca2+]i concentration to ~2.5 µM [4 ]. ATP evokes a similar rise in Ca2+ in mature BMDC, which exhibit little functional responses to caffeine or ryanodine (G. P. Ahern and P. J. O’Connell, unpublished observations) and in Ca2+-free medium [4 ], indicating that ATP-evoked Ca2+ transients occur predominantly via IP3-sensitive stores. Moreover, we show in this report that SP-evoked Ca2+ transients proceed normally during pharmacologic inhibition of RyRs with dantrolene. Thus, IP3Rs alone may be sufficient to elicit large Ca2+ increases in DC without the need for amplification by RyRs. These data suggest that RyRs in DC are not functionally coupled to IP3Rs and indeed may be activated by discrete signaling pathways. Recently, the subcellular localization of RyR was found to differ from IP3R in leukocytes [38 ]. Moreover, the subcellular localization of both Ca2+ mobilizing receptors is distinct among leukocyte subclasses, lymphocytes, monocytes, and neutrophils. The authors hypothesized that differential receptor trafficking may underlie their capacity for signaling a plethora of immune functions [38 ]. Similarly, a broad, intracellular distribution of IP3Rs may enable DC to maintain immune competence in the absence of RyR1 signaling. In summary, our data suggest that RyR1 is not obligate for normal DC development or for the ability of DC to take up antigen or stimulate T cells. We show that DC express all three isoforms of IP3R, suggesting a degree of functional redundancy in DC Ca2+ signaling pathways.


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ACKNOWLEDGEMENTS
 
This study was supported by a grant from the National Institutes of Health (NIAID AI055450) to G. P. A. and P. J. O. M. L-P. was supported by the Premiers’ Research Excellence Award (PREA; P. J. O.). We thank Dr. Paul D. Allen (PO1-AR17605) for the gift of RyR1 mutant mice and his critical review of this manuscript and Dr. Sean Cregan for advice about qPCR. We appreciate the assistance of Dägna Solveig Sheerar for cell sorting and Xiangbin Wang and Sandeep Pingle for cell culture.


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FOOTNOTES
 
2 Correspondence: Robarts Research Institute, 100 Perth Drive, London, ON, N6A 5K8, Canada. E-mail: peta{at}robarts.ca Back

Received December 16, 2005; revised May 10, 2006; accepted May 11, 2006.


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REFERENCES
 
    1
  1. Baixeras, E., Kroemer, G., Cuende, E., Marquez, C., Bosca, L., Ales Martinez, J. E., Martinez, C. (1993) Signal transduction pathways involved in B-cell induction Immunol. Rev. 132,5-47[CrossRef][Medline]
    2. 2
  2. Lewis, R. S., Cahalan, M. D. (1995) Potassium and calcium channels in lymphocytes Annu. Rev. Immunol. 13,623-653[Medline]
    3. 3
  3. Koski, G. K., Schwartz, G. N., Weng, D. E., Czerniecki, B. J., Carter, C., Gress, R. E., Cohen, P. A. (1999) Calcium mobilization in human myeloid cells results in acquisition of individual dendritic cell-like characteristics through discrete signaling pathways J. Immunol. 163,82-92[Abstract/Free Full Text]
    4. 4
  4. Hsu, S. F., O’Connell, P. J., Klyachko, V. A., Badminton, M. N., Thomson, A. W., Jackson, M. B., Clapham, D. E., Ahern, G. P. (2001) Fundamental Ca2+ signaling mechanisms in mouse dendritic cells: CRAC is the major Ca2+ entry pathway J. Immunol. 166,6126-6133[Abstract/Free Full Text]
    5. 5
  5. Czerniecki, B. J., Carter, C., Rivoltini, L., Koski, G. K., Kim, H. I., Weng, D. E., Roros, J. G., Hijazi, Y. M., Xu, S., Rosenberg, S. A., Cohen, P. A. (1997) Calcium ionophore-treated peripheral blood monocytes and dendritic cells rapidly display characteristics of activated dendritic cells J. Immunol. 159,3823-3837[Abstract]
    6. 6
  6. Rubartelli, A., Poggi, A., Zocchi, M. R. (1997) The selective engulfment of apoptotic bodies by dendritic cells is mediated by the {alpha}vß3 integrin and requires intracellular and extracellular calcium Eur. J. Immunol. 27,1893-1900[Medline]
    7. 7
  7. Gardella, S., Andrei, C., Lotti, L. V., Poggi, A., Torrisi, M. R., Zocchi, M. R., Rubartelli, A. (2001) CD8(+) T lymphocytes induce polarized exocytosis of secretory lysosomes by dendritic cells with release of interleukin-1ß and cathepsin D Blood 98,2152-2159[Abstract/Free Full Text]
    8. 8
  8. Gardella, S., Andrei, C., Poggi, A., Zocchi, M. R., Rubartelli, A. (2000) Control of interleukin-18 secretion by dendritic cells: role of calcium influxes FEBS Lett. 481,245-248[CrossRef][Medline]
    9. 9
  9. Andrei, C., Margiocco, P., Poggi, A., Lotti, L. V., Torrisi, M. R., Rubartelli, A. (2004) Phospholipases C and A2 control lysosome-mediated IL-1 ß secretion: implications for inflammatory processes Proc. Natl. Acad. Sci. USA 101,9745-9750[Abstract/Free Full Text]
    10. 10
  10. O’Connell, P. J., Wang, X., Leon-Ponte, M., Griffiths, C., Pingle, S. C., Ahern, G. P. (2006) A novel form of immune signaling revealed by transmission of the inflammatory mediator serotonin between dendritic cells and T cells Blood 107,1010-1017[Abstract/Free Full Text]
    11. 11
  11. Chan, V. W., Kothakota, S., Rohan, M. C., Panganiban-Lustan, L., Gardner, J. P., Wachowicz, M. S., Winter, J. A., Williams, L. T. (1999) Secondary lymphoid-tissue chemokine (SLC) is chemotactic for mature dendritic cells Blood 93,3610-3616[Abstract/Free Full Text]
    12. 12
  12. Montes, M., McIlroy, D., Hosmalin, A., Trautmann, A. (1999) Calcium responses elicited in human T cells and dendritic cells by cell-cell interaction and soluble ligands Int. Immunol. 11,561-568[Abstract/Free Full Text]
    13. 13
  13. Dieu, M. C., Vanbervliet, B., Vicari, A., Bridon, J. M., Oldham, E., Ait-Yahia, S., Briere, F., Zlotnik, A., Lebecque, S., Caux, C. (1998) Selective recruitment of immature and mature dendritic cells by distinct chemokines expressed in different anatomic sites J. Exp. Med. 188,373-386[Abstract/Free Full Text]
    14. 14
  14. Yanagihara, S., Komura, E., Nagafune, J., Watarai, H., Yamaguchi, Y. (1998) EBI1/CCR7 is a new member of dendritic cell chemokine receptor that is up-regulated upon maturation J. Immunol. 161,3096-3102[Abstract/Free Full Text]
    15. 15
  15. Delgado, E., Finkel, V., Baggiolini, M., Mackay, C. R., Steinman, R. M., Granelli-Piperno, A. (1998) Mature dendritic cells respond to SDF-1, but not to several ß-chemokines Immunobiology 198,490-500[Medline]
    16. 16
  16. Patterson, R. L., Boehning, D., Snyder, S. H. (2004) Inositol 1,4,5-trisphosphate receptors as signal integrators Annu. Rev. Biochem. 73,437-465[CrossRef][Medline]
    17. 17
  17. Meissner, G. (1994) Ryanodine receptor/Ca2+ release channels and their regulation by endogenous effectors Annu. Rev. Physiol. 56,485-508[CrossRef][Medline]
    18. 18
  18. Marks, A. R. (1997) Intracellular calcium-release channels: regulators of cell life and death Am. J. Physiol. 272,H597-H605[Medline]
    19. 19
  19. Franzini-Armstrong, C., Protasi, F. (1997) Ryanodine receptors of striated muscles: a complex channel capable of multiple interactions Physiol. Rev. 77,699-729[Abstract/Free Full Text]
    20. 20
  20. Hakamata, Y., Nishimura, S., Nakai, J., Nakashima, Y., Kita, T., Imoto, K. (1994) Involvement of the brain type of ryanodine receptor in T-cell proliferation FEBS Lett. 352,206-210[CrossRef][Medline]
    21. 21
  21. Guse, A. H., da Silva, C. P., Berg, I., Skapenko, A. L., Weber, K., Heyer, P., Hohenegger, M., Ashamu, G. A., Schulze-Koops, H., Potter, B. V., Mayr, G. W. (1999) Regulation of calcium signaling in T lymphocytes by the second messenger cyclic ADP-ribose Nature 398,70-73[CrossRef][Medline]
    22. 22
  22. Sei, Y., Gallagher, K. L., Basile, A. S. (1999) Skeletal muscle type ryanodine receptor is involved in calcium signaling in human B lymphocytes J. Biol. Chem. 274,5995-6002[Abstract/Free Full Text]
    23. 23
  23. Sei, Y., Gallagher, K. L., Daly, J. W. (2001) Multiple effects of caffeine on Ca2+ release and influx in human B lymphocytes Cell Calcium 29,149-160[CrossRef][Medline]
    24. 24
  24. O’Connell, P. J., Klyachko, V. A., Ahern, G. P. (2002) Identification of functional type 1 ryanodine receptors in mouse dendritic cells FEBS Lett. 512,67-70[CrossRef][Medline]
    25. 25
  25. Nakai, J., Dirksen, R. T., Nguyen, H. T., Pessah, I. N., Beam, K. G., Allen, P. D. (1996) Enhanced dihydropyridine receptor channel activity in the presence of ryanodine receptor Nature 380,72-75[CrossRef][Medline]
    26. 26
  26. Livak, K. J., Schmittgen, T. D. (2001) Analysis of relative gene expression data using real-time quantitative PCR and the 2(-{Delta} {Delta} C(T)) method Methods 25,402-408[CrossRef][Medline]
    27. 27
  27. Inaba, K., Inaba, M., Romani, N., Aya, H., Deguchi, M., Ikehara, S., Muramatsu, S., Steinman, R. M. (1992) Generation of large numbers of dendritic cells from mouse bone marrow cultures supplemented with granulocyte/macrophage colony-stimulating factor J. Exp. Med. 176,1693-1702[Abstract/Free Full Text]
    28. 28
  28. Christensen, J. L., Wright, D. E., Wagers, A. J., Weissman, I. L. (2004) Circulation and chemotaxis of fetal hematopoietic stem cells PLoS Biol. 2,E75[CrossRef][Medline]
    29. 29
  29. Sugiyama, T., Furuya, A., Monkawa, T., Yamamoto-Hino, M., Satoh, S., Ohmori, K., Miyawaki, A., Hanai, N., Mikoshiba, K., Hasegawa, M. (1994) Monoclonal antibodies distinctively recognizing the subtypes of inositol 1,4,5-trisphosphate receptor: application to the studies on inflammatory cells FEBS Lett. 354,149-154[CrossRef][Medline]
    30. 30
  30. Hirota, J., Baba, M., Matsumoto, M., Furuichi, T., Takatsu, K., Mikoshiba, K. (1998) T-cell-receptor signaling in inositol 1,4,5-trisphosphate receptor (IP3R) type-1-deficient mice: is IP3R type 1 essential for T-cell-receptor signaling? Biochem. J. 333,615-619[Medline]
    31. 31
  31. O’Connell, P. J., Pingle, S. C., Ahern, G. P. (2005) Dendritic cells do not transduce inflammatory stimuli via the capsaicin receptor TRPV1 FEBS Lett. 579,5135-5139[CrossRef][Medline]
    32. 32
  32. Hosoi, E., Nishizaki, C., Gallagher, K. L., Wyre, H. W., Matsuo, Y., Sei, Y. (2001) Expression of the ryanodine receptor isoforms in immune cells J. Immunol. 167,4887-4894[Abstract/Free Full Text]
    33. 33
  33. Bourguignon, L. Y., Chu, A., Jin, H., Brandt, N. R. (1995) Ryanodine receptor-ankyrin interaction regulates internal Ca2+ release in mouse T-lymphoma cells J. Biol. Chem. 270,17917-17922[Abstract/Free Full Text]
    34. 34
  34. Matsumoto, M., Nakagawa, T., Inoue, T., Nagata, E., Tanaka, K., Takano, H., Minowa, O., Kuno, J., Sakakibara, S., Yamada, M., Yoneshima, H., Miyawaki, A., Fukuuchi, Y., Furuichi, T., Okano, H., Mikoshiba, K., Noda, T. (1996) Ataxia and epileptic seizures in mice lacking type 1 inositol 1,4,5-trisphosphate receptor Nature 379,168-171[CrossRef][Medline]
    35. 35
  35. Futatsugi, A., Nakamura, T., Yamada, M. K., Ebisui, E., Nakamura, K., Uchida, K., Kitaguchi, T., Takahashi-Iwanaga, H., Noda, T., Aruga, J., Mikoshiba, K. (2005) IP3 receptor types 2 and 3 mediate exocrine secretion underlying energy metabolism Science 309,2232-2234[Abstract/Free Full Text]
    36. 36
  36. Sugawara, H., Kurosaki, M., Takata, M., Kurosaki, T. (1997) Genetic evidence for involvement of type 1, type 2 and type 3 inositol 1,4,5-trisphosphate receptors in signal transduction through the B-cell antigen receptor EMBO J. 16,3078-3088[CrossRef][Medline]
    37. 37
  37. Jayaraman, T., Ondriasova, E., Ondrias, K., Harnick, D. J., Marks, A. R. (1995) The inositol 1,4,5-trisphosphate receptor is essential for T-cell receptor signaling Proc. Natl. Acad. Sci. USA 92,6007-6011[Abstract/Free Full Text]
    38. 38
  38. Clark, A. J., Petty, H. R. (2005) Differential intracellular distributions of inositol trisphosphate and ryanodine receptors within and among hematopoietic cells J. Histochem. Cytochem. 53,913-916[Abstract/Free Full Text]



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L. Bracci, M. Vukcevic, G. Spagnoli, S. Ducreux, F. Zorzato, and S. Treves
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