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Injury and inflammation are potent regulators of adult neurogenesis. As the complement system forms a key immune pathway that may also exert critical functions in neural development and neurodegeneration, we asked if complement receptors regulate neurogenesis. We discovered that complement receptor 2 (CR2), classically known as a co-receptor of the B lymphocyte antigen receptor, is expressed in adult neural progenitor cells (NPCs) of the dentate gyrus. Two of its ligands, C3d and interferon-α (IFN-α), inhibited proliferation of wildtype NPCs but not NPCs derived from mice lacking Cr2 (Cr2−/−) indicating functional Cr2 expression. Young and old Cr2−/− mice exhibited prominent increases in basal neurogenesis compared with wildtype littermates, while intracerebral injection of C3d resulted in fewer proliferating neuroblasts in wildtype than in Cr2−/− mice. We conclude that Cr2 regulates hippocampal neurogenesis and propose that increased C3d and IFN-α production associated with brain injury or viral infections may inhibit neurogenesis.
The complement system is an important regulator of immune responses. Complement proteins can attract and activate various immune cells, amplify adaptive immune responses, promote phagocytosis, facilitate complement-mediated cytolysis by the membrane attack complex, and regulate cell proliferation and differentiation. Complement C3 is the central protein of the complement cascade, mediating its functions through various proteolytic fragments that bind to distinct complement receptors (CR) (Holers, 1996; Sahu and Lambris, 2001). Different C3 fragments can bind to CR1 (aka CD35), CR2 (CD21), CR3 (CD11b/CD18), CR4 (CD11c/CD18), as well as C3a receptor (C3aR) (Holers, 1996). Most of these receptors and their functions are conserved between mouse and human except CD35, which is an alternative splice product of the Cr2 gene in mouse but encoded by a separate gene in humans (Jacobson and Weis, 2008).
In the brain, complement can be produced by astrocytes, microglia, and neurons, and production is increased in brain injury and neurodegeneration (D'Ambrosio et al., 2001; Gasque, 2004). Genetic studies in mouse models have illustrated the complex function complement has in the brain. For example, overproduction of the rodent complement C3 inhibitor Crry (Wyss-Coray et al., 2002) or lack of C3 (Maier et al., 2008) led to increased accumulation of β-amyloid in mouse models for Alzheimer’s disease (AD), whereas lack of C1q reduced neurodegeneration in another AD mouse model (Fonseca et al., 2004). C1q appears to have a role in synaptic pruning during development (Stevens et al., 2007) and chronic increases of C1q observed in many neurodegenerative diseases have been postulated to promote synapse elimination. Interestingly, complement C3 and C3aR deficiency were associated with reduced neurogenesis in unmanipulated brains and after stroke (Rahpeymai et al., 2006) supporting a role for C3a, a potent inflammatory mediator, in the regulation of adult neurogenesis.
The adult mammalian brain can form new neurons in the subventricular zone of the lateral ventricles and the subgranular zone of the dentate gyrus in the hippocampus. Newly born neurons can integrate into functional circuits in the olfactory bulb or hippocampus, respectively. While the consequences of this process, particularly in the human brain, are unclear (Leuner et al., 2006; Zhao et al., 2008), hippocampal neurogenesis is required in forming certain types of memories in mice (Dupret et al., 2008; Zhang et al., 2008). Interestingly, injury stimulates (Dash et al., 2001), while inflammation inhibits neurogenesis (Monje et al., 2002; Ekdahl et al., 2003; Monje et al., 2003). Basal and injury-induced production of new hippocampal neurons dramatically decreases with age, as does the ability of neural progenitor cells to proliferate (Kuhn et al., 1996; Yagita et al., 2001; Jin et al., 2004). This may be the result of both, declining levels of mitogens, growth and survival factors (Anderson et al., 2002; Shetty et al., 2004) and increased abundance of inhibitory signals (Buckwalter et al., 2006) but more work is needed to understand the molecular mechanisms regulating adult neurogenesis. Because the complement system has broad functions in inflammation and cell homeostasis we asked whether complement receptors may regulate neurogenesis.
Mice deficient in the Cr2 gene (Cr2−/−) were obtained from Dr. Hector Molina, Washington University, MO (Molina et al., 1996). Mice deficient in the Cr3 gene (Cr3−/−) were obtained from Tanya Mayadas, Harvard Medical School, MA (Coxon et al., 1996). Human CR2/CD21 transgenic mice harboring a bacterial artificial chromosome BAC (hCD21 BAC; RP11-35C1; mouse line 38) express hCR2 under control of its natural human regulatory sequences (Hsiao et al., 2006). Cr2-Cre mice (B6.Cg-Tg(Cr2-cre)3Cgn/J; Stock No. 006368) harbor a transgene consisting of a cre recombinase expression cassette inserted into a BAC containing the mouse Cr2 locus (Kraus et al., 2004) and were obtained from The Jackson Laboratory (Bar Harbor, ME). We used two lines of Cre reporter mice: a double-fluorescent reporter mouse (B6.129(Cg)-Gt(ROSA)26Sortm4(ACTB-tdTomato,-EGFP)Luo/J; Stock No. 007676) which harbors a Chicken actin core promoter with a CMV enhancer driving membrane-targeted tandem dimer Tomato (mT)-polyA fragment flanked by loxP sites upstream of EGFP inserted into the ROSA26 locus (Muzumdar et al., 2007) or, a reporter mouse (B6. Cg-Gt(ROSA)26Sortm1(rtTA, EGFP) Nagy /J; Stock No. 005670) which harbors a PGK-neo-polyA fragment flanked by loxP sites upstream of rtTA and EGFP inserted into the ROSA26 locus (Belteki et al., 2005). Both mouse lines were obtained from The Jackson Laboratory and bred on an inbred, C57BL/6J genetic background using wildtype mice from The Jackson Laboratory. We used male and female mice in all experiments and did not observe any significant differences in the measurements reported herein. All animal care and use was in accordance with institutional guidelines and approved by the Veterans Administration Palo Alto Committee on Animal Research.
Immunostaining of mouse brain tissue was performed as described previously (Buckwalter et al., 2006). Briefly, brains of mice were immersed in 4% PFA overnight at 4°C, sunk through sucrose, and sectioned to 40 µm in thickness with a cryomicrotome. For 3,3′-Diaminobenzidine (DAB) staining, sections were incubated with primary antibodies rat anti-BrdU (0.4 µg/ml; Abcam) or goat anti-Dcx (0.4 µg/ml; Santa Cruz Biotechnology) overnight at 4°C, rinsed, and incubated with biotinylated rabbit anti-rat (1 µg/ml, vector) or rabbit anti-goat antibodies (3 µg/ml, (Vector Laboratories) followed by ABC labeling (Vector Laboratories) and development with DAB (Sigma). For BrdU labeling, brain sections were pre-treated with 2N HCl at 37°C for 30 min before incubation with primary antibody. For multi-label immunofluorescence of BrdU and cell type-specific markers, sections were incubated overnight with rat anti-BrdU, rinsed, and incubated for 1hr with donkey anti-rat antibody (2.5 µg/ml, Vector) before they were stained with one or two of the following antibodies: goat anti-Dcx, mouse anti-GFAP (1:1000; Chemicon International), mouse anti-NeuN (1.0 µg/ml, Chemicon International). To characterize GFP expressing cells in the spleen and the dentate gyrus of mT/mG mice, anti-Cr2 (clone 7E9, 1:100, Biolegend), anti-Sox2 (clone Y-17, 1:500, Santacruz Biotechnology) were used. Primary antibody staining was detected with secondary antibodies Cy3 donkey anti-rat (1:500, Jackson Immunoresearch), Alexa 488 donkey anti-goat (100 µg/ml, Invitrogen), Alexa 546 donkey anti-mouse (100 µg/ml, Invitrogen), Alexa 488 donkey anti-mouse (100 µg/ml, Invitrogen), or Alexa 488 donkey anti-rabbit (100 µg/ml, Invitrogen). Fluorescence images were obtained using a confocal-laser microscope (LSM 510 meta Pascal; Carl Zeiss MicroImaging, Inc.). Volocity software (Perkin Elmer) was used for 3D reconstruction with a coarse algorithm and deconvolution of Z-stack fluorescence images.
To quantify Dcx staining in 8 week-old mice, Metamorph imaging software (version 6.1r1; Universal Imaging Corporation) was used. The percentage of pixels above background staining within a region drawn around the dentate granule cell layer and subgranular zone was quantified. This method was used because the cell density was too high to count the cells in these 40 µm thick sections using light microscopy. In all other mice, Dcx positive cells in the same region were counted in every twelfth coronal section through the hippocampus and the total number of Dcx+ cells for each section was estimated by multiplying the number of cells counted by 12.
Fifty mg/kg of BrdU was injected intraperitoneally into 8-week-old mice once a day for 6 days, and mice were sacrificed 1 day (short-term labeling for figure 5) or 28 days (long-term labeling for figure 6) later. To estimate the total number of BrdU-positive cells in the brain, we performed DAB staining for BrdU on every sixth hemibrain section. BrdU+ cells in the granule cell and subgranular cell layer of the dentate gyrus in hemibrain were counted blinded with respect to genotype. The total number of BrdU+ cells in these hippocampal regions was then estimated by multiplying the number of cells counted by 12. Confocal microscopy was used to examine 50 to 100 BrdU-positive cells from each mouse for each stain to determine whether they co-labeled with Dcx, NeuN, or both, or with GFAP. The number of double- or triple-positive cells was expressed as a percentage of BrdU- positive cells.
Mouse neural progenitor cells were isolated from forebrains or hippocampi of P0–P3 mice and cultured as previously described (Reynolds and Rietze, 2005). Briefly, brains or hippocampi were dissected and incubated in DMEM containing 2.5 U/ml papain (Worthington Biochemicals), 250 U/ml DNaseI (Worthington Biochemicals) and 1 U/ml dispase II (Boeringher Mannheim) at 37°C for 45 min. Undigested pieces of tissue were removed by centrifugation at 500 g for 5 minutes and neural progenitor cells were isolated using 65% Percoll (Amersham Pharmacia Biotech) in DMEM with 10% FCS. Cellular pellets were resuspended in DMEM/F12 containing 5 mM HEPES buffer, 0.6% glucose, 3 mM sodium bicarbonate, 2 mM L-glutamine, 20 µg/ml insulin, 60 µM putrescine, 100 µṂ apotransferrin, 6.3 ng/mL progesterone, 5.2 ng/ml sodium selenite, 2 µg/ml heparin, 20 ng/mL EGF, and 10 ng/ml basic FGF, counted and plated in uncoated plates at 1 × 106 cells/ml. To prepare neurospheres from hippocampi, the percoll purification step was found not to be necessary and digested tissues were directly cultured in Neurobasal-A Medium (Invitrogen) supplemented with B27 (Invitrogen), Glutamax (Invitrogen), and the growth factors EGF (20 ng/ml; Peprotech) and bFGF (20 ng/ml; Peprotech). To passage cells, neurospheres were harvested by centrifugation (500 g for 5 minutes) and dissociated in Hanks-based cell dissociation buffer (Invitrogen). Rat neural progenitor cells were cultured as previously described (Monje et al., 2003). Briefly, cells were grown in DMEM/F12 supplemented with N2 supplement and 20 ng/ml basic FGF (Peprotech).
Total RNA was extracted using Trizol reagent (Invitrogen) and treated with TURBO DNA-free kit (Applied Biosystems) to digest DNA. Two microgram of DNase treated RNA samples were subjected to first-strand cDNA synthesis using oligo-dT primers supplied by the Taqman reverse transcription reagents kit (Applied Biosystems) by following the instruction manual. Primers used in this study are listed in supplementary table 1. For RT-PCR, GAPDH and GFP transcripts were amplified using ExTaq polymerase (Takara). Cr2 and Beta Actin transcripts were amplified using AmlpiTaqGold (Applied Biosystems) polymerase. For quantitative PCR, gene specific primer sets designed by either PrimerBank (http://pga.mgh.harvard.edu/primerbank/index.html) or RTPrimerDB (http://medgen.ugent.be/rtprimerdb/) were verified to produce single amplification products by gel electrophoresis. Using SYBR Green PCR Master mix (Applied Biosystems), reaction products were quantified and normalized to standard curves generated separately for each transcript from mRNA obtained from A20 cells (R2 = 0.9873 for Cr2, R2 = 0.9835 for GAPDH). SDS2.3 software from the ABI9700HT system (Applied Biosystems) was used to process data.
To assess the number of neurospheres 1 × 103 mouse NPCs at passage 5 were seeded per well in 96-well plates and incubated for 48 hrs at 37°C in the presence or absence of CR2 ligands and various antibodies. The total number of neurospheres in each well with diameters >50 µm were counted. Neurosphere size was evaluated using Metamorph imaging software (version 6.1r1; Universal Imaging Corporation). The cell number per dissociated sphere was analyzed from the same cultures using a standard Neubauer counting chamber. Cells were treated with the following proteins or antibodies: purified human C3 and human C3d (Complement Technology Inc.); recombinant mouse Interferon αA (IFNα, PBL Interferon Source); monoclonal rat anti-mouse CD35/CR2 antibody (clone 7G6; BD Biosciences); Polyclonal rabbit anti-human C3d (DAKO); biotin rat IgG2a isotype control antibody (Biolegend).
1 × 104 rat NPCs were incubated with C3 or C3d at the indicated concentration at 37°C for 48 hrs prior to cell counting. A rabbit antibody against human C3d (1:1000, DAKO) was used for blocking C3d.
Purified human C3d (0.5 µg in 0.5 µl) was injected stereotaxically into the right side of the dentate gyrus and an equal amount of saline was injected into the left side (1.75 mm posterior, 0.75 mm lateral, 1.7 mm ventral from bregma). After mice gained consciousness from surgery 50 mg/kg BrdU was injected intraperitoneally once and then again daily for 5 days until mice were sacrificed on day 7. Brains were removed, immersed in 4% PFA overnight at 4°C, sunk through sucrose, and sectioned to 40 µm in thickness with a cryomicrotome. Five sections close to the injection site were then stained with antibodies against BrdU and Dcx as described above. The number of BrdU+/Dcx+ double positive cells was counted using confocal microscopy on the C3d or saline injected sides on the one section most proximal to the injection site per mouse and the ratio of double labeled cells between C3d and saline injected sides was calculated.
All immunohistochemical experiments were analyzed by an investigator blinded to genotype or age. Values are presented as mean + SEM. Differences among groups were analyzed using t-test or one-way ANOVA with Scheffe post hoc test. P-values ≤ 0.05 were considered to be significant. Analyses were done using Statview Software.
Injury and inflammation are potent regulators of adult neurogenesis (Dash et al., 2001; Monje et al., 2002; Ekdahl et al., 2003; Monje et al., 2003). To determine if complement factors are involved in this process we asked whether complement receptors are expressed in neural progenitor cells (NPCs). While CR3 (CD11b/CD18) and CR4 (CD11c/CD18) are accepted markers for the myeloid lineage, expression of the Cr2 gene product CR2, which is involved in multiple stages of B lymphocyte differentiation and proliferation (Tolnay and Tsokos, 1998; Carroll, 2004) has not been studied in neurogenesis. To track Cr2 gene expression in vivo we crossed Cr2-cre transgenic mice (B6.Cg-Tg(Cr2-cre)3Cgn/J) (Kraus et al., 2004), which harbor cre recombinase inserted into a mouse Cr2 BAC clone, with ROSA26-Actin promoter-loxP-mT-pA-loxP-mEGFP-pA reporter mice (herein called mT/mG mice) (Muzumdar et al., 2007). In double transgenic mice cre-induced recombination in Cr2 expressing cells leads to excision of the gene encoding for membrane targeted tandem dimer Tomato (mT) and a stop sequence resulting in turn in expression of membrane targeted EGFP. Thus, every cell in this mouse either expresses Tomato or EGFP. To validate the Cr2-driven Cre reporter system, we stained spleen tissue sections with the anti-Cr2 antibody 7E9 and compared it with Cr2-cre induced EGFP protein expression in 2-month-old Cr2-cre;mT/mG reporter mice. We observed that all EGFP expressing cells also labeled with the Cr2 antibody (Fig. 1A). In the brain, EGFP positive cells were found throughout the hippocampus (Fig. 1B) suggesting that these cells are still expressing CR2 or that they descended from CR2 expressing precursors. Higher power confocal images showed that many of the cells in the subgranular zone of the dentate gyrus (SGZ) also expressed Sox2 consistent with characteristics of neural progenitor cells (Fig. 1C, D, Supporting Video 1).
Because none of the available Cr2 antibodies produced specific staining on brain sections if Cr2 knockout mice were used as a control (Supplemental Table 2) we measured Cr2 mRNA in brain tissue and cells. In support of Cr2 promoter activity in NPCs, EGFP mRNA was expressed in freshly isolated neonatal NPCs from Cr2-cre;ROSA26-GFP reporter mice which harbor ROSA26-loxP-stop-loxP-rtTA-IRES-EGFP reporter construct (Fig. 2A). Additional evidence that the CR2 promoter is active in NPCs was obtained from “humanized” CR2 transgenic mice in which an artificial bacterial chromosome containing the human CR2 locus (which does not contain the independent CD35 gene) leads to human CR2 (hCD21) expression under control of the human promoter, for example, in B cells (Hsiao et al., 2006). In addition, the mice were crossed onto the Cr2−/− background, essentially replacing mouse Cr2 with human CR2. Notably, CD35 (endogenous mouse and human) is lacking in these mice. We observed that NPCs from these hCD21-BAC;Cr2−/− mice expressed human CR2 mRNA whereas NPCs from nontransgenic mice did not (Fig. 2B). These findings strongly suggest that NPCs express CR2. Lastly, neonatal wildtype primary mouse NPCs expressed Cr2 mRNA by RT-PCR whereas Cr2−/− NPCs did not (Fig. 2C). To assess relative levels of Cr2 expression, we compared transcript levels in NPCs with those in lymphoid tissue and other CNS cells using quantitative PCR (qPCR). Although NPCs isolated from C57Bl/6 or FVB mouse strains expressed less than 1% of Cr2 mRNA compared with spleen or B cell line A20, they expressed 3–4 times the amount detected in primary astrocytes (Fig. 2D). Microglia showed no detectable Cr2 mRNA (Fig. 2D). Because CD35 and CR2 are splice products encoded by the same gene in mice (i.e. Cr2) and differ from each other only by several identical so-called short consensus repeats (Carroll, 2004) we were not able to amplify specific transcripts discerning the splice variants.
CR2 is a key module of the B cell antigen receptor complex together with CD19, CD81, and CD225, and therefore we asked whether these other components are also expressed in NPCs. Indeed, we were able to detect mRNAs encoding for all three components (Fig. 1S), which are involved in part in B cell receptor signal transduction. Together, these findings show Cr2 is expressed in mouse NPCs and raise the question whether it is functionally active.
In B cells, CR2 together with several adaptor proteins functions as the co-receptor for membrane-bound immunoglobulins (i.e. the B cell receptor) by binding antigens coated with the complement adjuvant C3d, thus playing an important role in the activation and proliferation of B cells (Tolnay and Tsokos, 1998; Carroll, 2004). To determine the functionality of CR2 in NPCs, we cultured primary postnatal NPCs derived from hippocampi or the entire forebrain from Cr2−/− mice (Molina et al., 1996) and wildtype littermate controls, as well as previously characterized rat NPC lines (Monje et al., 2003) in the presence and absence of CR2 ligands. We first tested the effect of human C3d, which is a functional ligand of the mouse CR2 receptor (Molina et al., 1991). In the absence of ligands there was no consistent difference in the number or size of neurospheres at various passages between cultures derived from Cr2−/− mice or wildtype littermates (Fig. S2), and there were no apparent morphological differences in cell size or shape (data not shown). Treatment of hippocampal derived wildtype primary mouse NPCs with purified human C3d reduced neurosphere number by a third but had no effect on Cr2−/− NPCs (Fig. 3a). Consistent with these findings, C3d also reduced proliferation in the rat NPC line, while C3d incubated with C3d neutralizing antibody or intact C3 had no effect on cell proliferation (Fig. 3B). Complement C3d or C3 did not induce overt cell death as no significant increase in lactate dehydrogenase (LDH) could be measured in culture supernatants from C3d or C3 treated compared with untreated NPCs (data not shown). The cytokine IFN-α has recently been shown to signal through the same domains of human CR2 as C3d, and to bind to it with similar affinity (Asokan et al., 2006). Consistent with being a ligand for CR2, IFN-α significantly inhibited proliferation in wildtype but not Cr2−/− primary mouse NPCs (Fig. 3A). Furthermore, the decrease in proliferation seen in wildtype primary mouse NPCs was reversed by the addition of antibody 7G6, which blocks ligand binding to CR2 (Martin et al., 1991), further supporting that C3d and IFN-α inhibit NPC proliferation in our cell culture system by binding to CR2. In line with these results, neurosphere number and the number of cells per neurospheres were reduced following C3d or IFN-α treatment in wildtype NPCs derived from entire mouse forebrains but not in Cr2−/− NPCs (Fig. S3). In summary, these findings demonstrate the functional expression of Cr2 in cultured mouse and rat NPCs and show that CR2 ligands inhibit NPC proliferation.
To investigate whether CR2/CD35 have a role in adult neurogenesis in vivo we assessed expression of doublecortin (Dcx), a marker of proliferating neuroblasts and immature neurons, in Cr2−/− and wildtype littermates. We also analyzed neurogenesis in mice deficient in Cr3, a major phagocyte receptor restricted to microglia and macrophages in the brain, or in crosses resulting from these mice and lacking both, Cr2 and Cr3. Lack of Cr2 resulted in a two-fold increase in Dcx immunoreactivity in the dentate gyrus at 8 weeks and 5–7 months of age (Fig. 4A–C), and an almost 3-fold increase in the number of Dcx+ cells in 13–16-month-old Cr2−/− mice compared to wildtype controls (Fig. 4A,D). In contrast, littermate mice deficient in Cr3 showed no changes in the number of Dcx expressing cells compared to wildtype controls (Fig. 4D). Mice lacking Cr2 and Cr3 showed equal increases in Dcx expressing cells as Cr2−/− mice (Fig. 4D).
To determine whether lack of Cr2 led to an increase in proliferation in the hippocampus, the nucleotide analog BrdU was injected into mice for 6 days and the commitment of newly generated cells was evaluated 1 day later. Neural progenitor cells in the subgranular zone of the dentate gyrus are capable of differentiating into either the neuronal or glial lineage. While the total number of BrdU+ cells in the dentate gyrus was not significantly different between wildtype and Cr2−/− mice in this short term labeling paradigm (Fig. 5A), the number of BrdU+/Dcx+ cells was slightly but significantly increased in Cr2−/− mice (Fig. 5B–D). As expected, over this short time only few dividing cells acquired a NeuN positive phenotype and this was not changed by the lack of Cr2. The commitment of NPCs to the glial lineage was assessed using GFAP as a marker for astrocytes. No significant difference was observed in BrdU+/GFAP+ cells between wildtype and Cr2−/− mice following short term labeling with BrdU (Fig. 5B–D). Nevertheless, out of all dividing cells the percentage of GFAP expressing and other non-neuronal dividing cells was reduced from 32.6% to 20.8% in Cr2−/− mice compared with wildtype littermate controls while the number of cells in the neuronal lineage increased from 67.4% to 79.2% (Fig. 5D).
To determine if the increase in Dcx+ cells observed in Cr2−/− mice is the result of changes in the maturation and survival of newly generated immature neurons we analyzed the number of label retaining neurons 28 days after BrdU injection. In this long term labeling paradigm Cr2−/− mice at 8 weeks of age (Fig. 6A), and more profoundly, at 5–7 months of age (Fig. 6A) had significantly more BrdU+ cells overall than wildtype mice and there were more mature BrdU/NeuN+ neurons in Cr2−/− mice at 8 weeks of age (Fig. 6B,C).
To determine whether Cr2 encodes a functional gene product in NPCs and CR2/CD35 signaling affects neurogenesis in vivo we stereotaxically injected C3d into the right dentate gyrus of wildtype or Cr2−/− mice. Saline was injected as a control into the left dentate gyrus and BrdU was given for 6 days intraperitoneally until the mice were sacrificed the next day. Injection of C3d resulted on average in a 35% decrease in the number of BrdU/Dcx+ cells in the ipsilateral compared with the saline injected side. Consistent with cell culture data, C3d had no effect in Cr2−/− mice (Fig. 7). These findings further corroborate the cell culture studies suggesting that CR2/CD35 is functionally expressed in the hippocampus and can modulate adult neurogenesis in vivo.
The complement system is best known for its critical functions in the immune system but a growing number of studies have tied complement factors genetically to neurodegenerative diseases in humans and mice (Wyss-Coray et al., 2002; Haines et al., 2005; Klein et al., 2005; Maier et al., 2008). Most recently, the human CR1 locus was linked to AD in a large genome wide association study (Lambert et al., 2009), although the biological role of CR1 in that disease, or in the brain in general, is unknown. Here we describe that Cr2, best known for encoding a key component of the B cell receptor complex, is functionally expressed in NPCs. We demonstrate that Cr2−/− mice exhibit increased numbers of immature and mature neurons in the hippocampus implicating Cr2 in the maintenance of adult neurogenesis. Conversely, C3d and IFN-α, two independent ligands of human CR2/CD21, inhibit NPC proliferation in cell culture and injection of C3d into the dentate gyrus reduces the number of immature neuroblasts in vivo.
Our findings add C3d and IFN-α to a growing number of immune molecules and inflammatory proteins that regulate adult neurogenesis in rodents. For example, interleukin-1β (Koo and Duman, 2008), interleukin-6 (Monje et al., 2003), tumor necrosis factor (TNF)-α (Iosif et al., 2006), and transforming growth factor-beta (Buckwalter et al., 2006; Wachs et al., 2006) inhibit neurogenesis. It is unclear if these factors are merely sufficient to inhibit adult neurogenesis when increased, or whether they are also required to exert a certain negative signal to maintain homeostatic levels of neurogenesis. On the other hand, immune regulatory factors such as IL-6, leukemia inhibitory factor (LIF), or cliliary neurotrophic factor that signal through gp130 can stimulate neural stem cell proliferation or maintain stem cell fate (Shimazaki et al., 2001; Bauer and Patterson, 2006). The relative importance of these proteins needs to be determined as it is unclear whether they act alone or in concert, and whether individual proteins are restricted to specific injury and disease conditions.
We report that lack of Cr2 in mice leads to a 2–3-fold increase in the number of immature neuroblasts (Figure 4) and a roughly 40% increase in mature neurons (Figure 6C) in the young adult hippocampus, suggesting CR2/CD35 may have a physiological role in the control of NPC number in the adult mouse by exerting a tonic inhibitory signal. In support of this, lineage tracing studies using a Cr2-cre controlled reporter gene indicate that Cr2 is expressed in a small set of cells in the subgranular layer of the dentate gyrus that show neural stem cell-like characteristics expressing both Sox2 (Fig. 1). Lack of Cr2 may also lead to an expansion of such early, possibly uncommitted cells but more studies are needed to prove this point. Low level Cr2 reporter gene expression was also noted in dentate gyrus granule neurons and very low levels of Cr2 transcripts were detected in primary astrocytes (Fig. 2D). CR2 expression had previously been documented in human glioma cells and fetal astrocytes (Gasque et al., 1996) but its expression in mice seems much lower.
We do not know how Cr2 deficiency causes the observed increase in neurogenesis and which splice form of the Cr2 gene mediates this effect. Mouse CD35 and CR2 are spliced from the same gene and no conditions have been reported to favor splicing of one or the other product, suggesting both isoforms are expressed on NPCs similar to their co-expression on B cells (Jacobson and Weis, 2008). Furthermore, it is predicted that mouse CR2 ligands would also bind to mouse CD35 making it difficult to exclude a role for CD35 in the observed effects (Molina et al., 1994). Additional studies will also be necessary to address whether CR2 on NPCs forms a similar signaling complex as the one found in B cells with CD19, CD81, and CD225. Notably, all three genes seem to be expressed in NPCs (Fig. S1).
Based on our results in CR2/CD35 deficient mice physiological levels of endogenous ligands may function to control basal neurogenesis whereas increased levels of complement C3d or IFN-α following injury may lead to detrimental inhibition of neurogenesis. Human CR2 is the main receptor for C3d; it is also a receptor for IFN-α (although IFN-α signals through classical IFN-α receptors as well), CD23, and the EBV envelope protein gp350 (Asokan et al., 2006). Interestingly, basal neurogenesis in the dentate gyrus was significantly reduced not only in C3aR knockout mice but also in C3 knockout mice (Rahpeymai et al., 2006), which cannot generate the C3 cleavage product C3a. Because C3d is also derived from C3 we would expect neurogenesis to increase in C3 knockout mice if C3d signaling was actively suppressing neurogenesis via Cr2. We therefore think C3d is not responsible for suppressing neurogenesis via Cr2 under physiological conditions. Nevertheless, C3d appears to be sufficient to reduce the number of immature neuroblasts in vivo (Fig. 7), and may exert such a function following CNS injury. We think it is more likely that CR2 ligands exert their effect by reducing proliferation of neuroblasts rather than by increasing apoptosis because we were unable to detect an increase in activated caspase staining in vivo, though the methods successfully detected alcohol induced cell death (data not shown). With respect to IFN-α, at least 13 functional subtypes have been described in humans (Theofilopoulos et al., 2005) and it is unclear whether they are constitutively produced at levels that might trigger CR2 signaling in NPCs in the normal brain and whether they differ in their affinity for CR2. Even without knowing the key endogenous ligand therapeutic inhibition of CR2 signaling may promote basal neurogenesis in aging or restore neurogenesis in a number of inflammatory conditions where C3d or IFN-α are elevated and neurogenesis is reduced.
Adult hippocampal neurogenesis is being linked ever more strongly to the formation of specific new memories in rodents (Dupret et al., 2008; Zhang et al., 2008) and it is possible that neurogenesis in the human hippocampus may have a similar role (Leuner et al., 2006; Zhao et al., 2008). Although this issue is far from being settled, our findings may shed new light on an intriguing association of increased levels of C3d or IFN-α with memory problems or neuropsychiatric conditions in several human diseases. For example, C3d levels are increased in HIV (Fust et al., 1991), and increased levels of IFN-α in the CSF have been linked with HIV dementia (Rho et al., 1995). Similarly, C3d (Rother et al., 1993) and IFN-α (Harrison and Ravdin, 2002; Schmidt and Ouyang, 2004) levels are increased in autoimmune diseases such as systemic lupus erythematosus (SLE), which frequently presents with cognitive impairment. Perhaps the strongest evidence for a possible link between IFN-α and cognition comes from patients treated with recombinant IFN-α as part of a therapy for hepatitis C virus infection or cancer (Pavol et al., 1995; Scheibel et al., 2004; Hilsabeck et al., 2005). At least one third of patients have been reported to develop depression or cognitive deficits linked to the treatment, and normal function is regained after treatment is discontinued. Consistent with these observations in humans adult rats treated with IFN-α showed depression-like symptoms and reduced cell proliferation (BrdU incorporation) in the dentate gyrus (Kaneko et al., 2006). While these authors had no mechanistic explanation of their findings our study suggests that the observed effects of IFN-α in rats and possibly in humans could be mediated through CR2 expressed on NPCs. Together with published studies, our findings support a scenario where abnormal CR2 signaling in certain viral infections and autoimmune disorders inhibits neurogenesis and as a result leads to depressive symptoms and cognitive impairments.
This work was supported by a postdoctoral fellowship from the Japanese Society for Promotion of Sciences (M.M.), and the National Institute on Aging (AG20603 and AG27505 to T.W.-C.). We thank Dr. T. Palmer for the rat NPC line and advice on the culture and analysis of NPCs, Dr. M. Buckwalter for technical advice, and Y. Takeda-Uchimura for technical support.
CONFLICT OF INTEREST: none
The Supplemental Data for this article can be found online.