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Originally published In Press as doi:10.1074/jbc.M604959200 on August 2, 2006

J. Biol. Chem., Vol. 281, Issue 39, 29076-29084, September 29, 2006
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Transgenic Mice Expressing Dominant-negative Activin Receptor IB in Forebrain Neurons Reveal Novel Functions of Activin at Glutamatergic Synapses*

Mischa Roland Müller{ddagger}, Fang Zheng§, Sabine Werner{ddagger}12, and Christian Alzheimer§13

From the {ddagger}Institute of Cell Biology, Department of Biology, ETH Zurich, CH-8093 Zurich, Switzerland and the §Institute of Physiology, University of Kiel, Olshausenstrasse 40, D-24098 Kiel, Germany

Received for publication, May 23, 2006 , and in revised form, July 25, 2006.


   ABSTRACT
TOP
 ABSTRACT
INTRODUCTION
 EXPERIMENTAL PROCEDURES
 RESULTS
 DISCUSSION
 REFERENCES
 
The transforming growth factor beta family member activin is an important regulator of development and tissue repair. It is strongly up-regulated after acute injury to the adult brain, and application of exogenous activin protects neurons in several lesion models. To explore the role of endogenous activin in the normal and acutely damaged brain, we generated transgenic mice expressing a dominant-negative activin receptor IB (dnActRIB) mutant in forebrain neurons. The functionality of the transgene was verified in vivo. Hippocampal neurons from dnActRIB mice were significantly more vulnerable to intracerebroventricular injection of the excitotoxin kainic acid than those from control littermates, indicating a crucial role of endogenous activin in the rescue of neurons from excitotoxic insult. Because dnActRIB is only expressed in neurons, but not in glial cells, activin affords protection at least in part through a direct action on endangered neurons. Unexpectedly, the transgenic mice also revealed a prominent novel role of activin in glutamatergic neurotransmission in the intact adult brain. Electrophysiologic examination of excitatory synapses onto CA1 pyramidal cells in hippocampal slices of dnActRIB mice showed a reduced NMDA current response, which was associated with impaired long term potentiation. This is the first demonstration that activin receptor signaling is essential to optimize the performance of neuronal circuits in the mature brain under physiological conditions.


   INTRODUCTION
TOP
 ABSTRACT
 INTRODUCTION
EXPERIMENTAL PROCEDURES
 RESULTS
 DISCUSSION
 REFERENCES
 
Activins are members of the transforming growth factor beta family of proteins, which regulate proliferation and differentiation of various cell types. The predominant activin variants are homo- or heterodimers consisting of betaA and/or betaB subunits (activin A: betaAbetaA, activin B: betaBbetaB and activin AB: betaAbetaB). Their biological effects are mediated through heteromeric receptor complexes, consisting of type I and type II transmembrane serine/threonine kinase receptors (1). Upon ligand binding, one of the activin type II receptors (ActRII or ActRIIB) dimerizes with a type I receptor (ActRIA, ActRIB, or activin receptor-like kinase 7 (2, 3)), resulting in phosphorylation of the type I receptor by the type II receptor. The subsequently activated serine/threonine kinase of the type I receptor phosphorylates the recruited receptor Smads (Smad2 and Smad3). The latter translocate to the nucleus upon multimerization with Smad4 and modulate as transcription factor complexes the expression of activin target genes (4). In addition, other signaling pathways are also activated by activin receptors, including mitogen activated kinase signaling (1, 4, 5).

Activins are important regulators of development, inflammation, and repair of different tissues and organs (1, 5). In the central nervous system, activin is involved in development, but it may also participate in adaptive and protective mechanisms of the adult brain. Thus, activin betaA mRNA is transiently up-regulated after electrical stimuli that induce synaptic long term potentiation (LTP4; Refs. 6 and 7) and after brief electroconvulsive seizures that are used to treat certain forms of major depression (8). A much stronger and prolonged induction of activin A expression occurs after acute brain injury (9-11). Since exogenous activin protects cultured neurons (12, 13), these findings suggested that activin might also afford neuroprotection in vivo. Indeed, intracerebroventricular application of activin A rescued neurons after excitotoxic and hypoxic/ischemic insults (10, 14, 15). Nevertheless, the site of action and the function of endogenous activin in the intact and the injured brain remain to be determined. Therefore, we generated transgenic mice expressing a dominant-negative mutant of activin receptor IB (dnActRIB) in forebrain neurons. For this purpose, the complete kinase domain of the receptor was deleted. When expressed in excess in comparison to the endogenous receptor, dnActRIB preferentially forms non-functional heterodimers with type II receptors upon ligand binding, thereby blocking activin receptor signaling (16). Using this strategy, we report here the novel and unexpected finding that activin signaling augments plasticity of excitatory synapses in the adult brain, while, at the same time, reducing its vulnerability to excitotoxic damage.


   EXPERIMENTAL PROCEDURES
TOP
 ABSTRACT
 INTRODUCTION
 EXPERIMENTAL PROCEDURES
RESULTS
 DISCUSSION
 REFERENCES
 
Generation of Transgenic Mice—The transgene construct (Fig. 1A) was separated from vector sequences (Fig. 1A), and the purified insert was used for microinjection of fertilized oocytes from B6D2F2 x C57BL/6 mice. Mice were genotyped by PCR, and transgenic animals were backcrossed into the C57BL/6 strain.

Intracerebroventricular Injection of KA, Activin A, or PBS— Mice were anesthetized with ketamine/xylazine and placed into a stereotaxic apparatus. Intracerebroventricular injection of 0.1 µlofKA(1 µg/µl in PBS, pH 7.0) was performed as described by Tretter et al. (9, 10) with permission from the veterinary authorities, Zurich, Switzerland. To verify the functionality of the transgene, 150 ng of activin A in 0.8 µl of PBS or solvent PBS were injected intracerebroventricularly. Ipsilateral hippocampi were removed 2 h later and used for preparation of protein lysates and subsequent Western blotting.

RNase Protection Assay (RPA)—RNA was isolated from hippocampi, and RPAs were performed (9) using a 507-bp probe corresponding to the 5'-end of the murine ActRIB cDNA and some region of the hybrid intron present in the expression vector (Fig. 1A) and a fragment corresponding to nucleotides 566-685 of the murine glyceraldehyde-3-phosphate dehydrogenase cDNA.

In Situ Hybridization—In situ hybridization with a digoxygenin-labeled riboprobe was performed on frozen sections from whole brains as described by Yang et al. (17) using a riboprobe corresponding to sequences of the polyadenylation signal of the vector (Fig. 1A).

Western Blotting—Hippocampi were lysed in 50 mM Tris/HCl, pH 7.4, 150 mM NaCl, 1 mM EDTA, 1% Nonidet P-40, 0.25% sodium deoxycholate, 10 µg/ml each of aprotinin, leupeptin, and pepstatin, 0.5 mM AEBSF, 10 mM NaF, 1 mM Na3VO4, 10 mM sodium pyrophosphate, 20 µM phenylarsinoxide, phosphatase inhibitor mixture I/II (Sigma, Munich, Germany; 1:100). Lysates were analyzed by Western blotting using polyclonal antibodies against the c-Myc epitope, total Smad2 and Smad3 (both from Santa Cruz Biotechnology, Santa Cruz, CA), or phospho-Smad2 (pSmad2; kindly provided by Dr. C.-H. Heldin, Uppsala, Sweden). In addition, a monoclonal antibody against beta-actin (Sigma) was used.

Immunostaining—Frozen sections (25 µm) from normal and injured hippocampus were fixed with ice-cold methanol, and endogenous peroxidase activity was quenched by a 20 min treatment with 0.3% H2O2 in methanol. Sections were then treated for 30 min with PBS, 3% BSA, 0.02% Nonidet P-40 to block unspecific binding sites and subsequently incubated with a rabbit polyclonal antibody against cleaved caspase-3 (Asp175, Cell Signaling Technology Inc., Danvers, MA). After incubation with the secondary antibody (anti-rabbit IgG, peroxidase-coupled (Promega, Wallisellen, Switzerland), they were stained with diaminobenzidine in 0.01% H2O2.

For immunofluorescence analysis, frozen sections were fixed with ice-cold methanol and unspecific binding sites were blocked as described above. Sections were incubated with a rabbit polyclonal antibody against phospho-Smad2 (see above), and phospho-Smad2-positive cells were detected with Cy3-labeled anti-rabbit IgG (Jackson Immunoresearch, Westgrove, PA).

Real-time RT-PCR—RNA from total hippocampi was reverse transcribed and analyzed by quantitative real-time PCR using SYBR Green technology as described by the manufacturer (Applied Biosystems, Rotkreuz, Switzerland). RNA samples, which had not been reversely transcribed were used as a negative control. The following primers were used: NMDA receptor 1 (NR1), 5'-GCC CAA CGC CAT ACA GAT G-3' and 5'-GGC GGG TGA CTA ACT AGG ATA GC-3'; NR2A, 5'-TGC TAC GGG CAG ACA GAG AAG-3' and 5'-GCC ATC CCA AGT CAC ATT GAC-3'; NR2B, 5'-GAC CTG CAT GCG GAA TAC AGT-3' and 5'-TAG CCT GGT TCC TCA TCT GTT TTA-3'; beta-actin, 5'-CGT GAA AAG ATG ACC CAG ATC A-3' and 5'-CAC AGC CTG GAT GGC TAC GT-3'.

Quantification of Neuronal Survival—Frozen coronal brain sections (25 µm) were stained with cresyl violet (Nissl staining) or Fluoro-Jade B (FJB) (Histo-Chem. Inc, Jefferson, AR; 0.0004% in 0.1% acetic acid) as described by Schmued and Hopkins (18). FJB-stained sections were analyzed using ImageJ software (W. S. Rasband, National Institutes of Health, Bethesda, MA). After background subtraction, inversion, and calibration of the picture, a binary was made, and the area of all black pixels was measured. Statistical analysis was performed using the Mann-Whitney U test for non-Gaussian distribution.

Preparation of Hippocampal Slices and Electrophysiology— Transverse hippocampal slices (350 µm) were prepared from the brain of control and mutant mice (2-4 months of age). Slices were constantly superfused with artificial cerebrospinal fluid composed of (in mM) 130 NaCl, 3 KCl, 2 CaCl2, 2 MgCl2, 1.25 NaH2PO4, 25 NaHCO3, and 10 D-glucose, gassed with 95% O2, 5% CO2, pH 7.4. Field excitatory postsynaptic potential (fEPSP) recordings from stratum radiatum of area CA1 were performed and analyzed by means of a multielectrode system (MED64 in conjunction with Performer 2.0 software, Panasonic, Japan). In the recording chamber, submerged slices were constantly superfused with warmed artificial cerebrospinal fluid (30 °C). Before and after tetanization, test and control pathways were stimulated at 0.05 Hz. The stimulus strength was adjusted to obtain ~40% of the maximum fEPSP amplitude under control conditions. LTP was induced by two theta bursts stimuli (TBS) 20 s apart, with each TBS consisting of 15 bursts of 4 pulses at 100 Hz delivered at an interburst interval of 200 ms. fEPSPs slopes were normalized to 100% before TBS and pooled across animals of the same genotype. Data were processed and analyzed using Origin Pro7 software (OriginLab, Northampton, MA) and were given as mean ± S.E. Pipettes for whole-cell recordings of excitatory postsynaptic currents (EPSCs) were filled with (in mM) 130 potassium gluconate, 3 MgCl2, 5 EGTA, 5 HEPES, 2 Na2-ATP, 0.3 Na-GTP, pH 7.25-7.30. Series resistance (10-20 M{Omega}) was compensated by 70-85%. EPSCs were recorded in the presence of the GABAA ({gamma}-aminobutyric acid, type A) receptor antagonist bicuculline (30 µM) at -70 mV, after correcting for liquid junction potentials. Whole-cell recordings were performed at room temperature. Current signals were recorded and analyzed using an Axopatch 200 amplifier in conjunction with a Digidata 1200 interface and pClamp 9 software (all from Axon Instruments). Data were expressed as means ± S.E. Statistical comparisons were performed using Student's t test. All drugs were from Sigma.


Figure 1
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  		FIGURE 1.
Generation of dnActRIB transgenic mice. A, transgene construct, including the CaMKII-{alpha} promoter, a hybrid intron consisting of an adenovirus splice donor site and an IgG splice acceptor site, the dnAc-tRIB cDNA fused to the coding sequence of the c-Myc epitope, and the SV40 polyadenylation site. ECD, extracellular domain; TMD, transmembrane domain. Riboprobes used for RPA/in situ hybridization are indicated below. B,10 µg of total RNA from hippocampi of wt and tg mice of different lines (L1, L2, and L3) were analyzed by RPA for the presence of endogenous (end.) ActRIB mRNA or transgene-derived (tg dnActRIB) mRNA or contaminating ActRIB DNA (DNA). 10 µg of tRNA or 10 ng of the transgene plasmid were used as negative or positive controls, respectively. 1000 cpm of the hybridization probes were loaded in the lanes labeled "probes." Hybridization with a glyceraldehyde-3-phosphate dehydrogenase (gapdh) riboprobe served as a loading control. C, lysates from hippocampi of wild-type and transgenic mice were analyzed for the presence of dnActRIB and of beta-actin. D-F, frozen brain sections from wild-type (F) and transgenic mice (D and E) were hybridized with antisense (D and F) and sense (E) probes corresponding to a region specific for the transgene-derived mRNA. Bar = 100 µm.

 

   RESULTS
TOP
 ABSTRACT
 INTRODUCTION
 EXPERIMENTAL PROCEDURES
 RESULTS
DISCUSSION
 REFERENCES
 
Generation of Transgenic Mice Expressing dnActRIB in Hippocampal Neurons—To inhibit activin receptor signaling in hippocampal neurons in vivo, we generated transgenic mice expressing dnActRIB with a carboxyterminal c-Myc epitope under the control of the CaMKII-{alpha} promoter and regulatory elements (19) (Fig. 1A). The progeny of seven transgenic founder mice expressed the transgene as determined by RPA using a riboprobe, which can distinguish between transgene-derived and endogenous ActRIB mRNA (Fig. 1B and data not shown). Lines 1 and 3 with the highest expression level were used for further studies. The transgene-derived protein was detected by Western blotting using an antibody against the c-Myc epitope (Fig. 1C). In situ hybridization with a transgene-specific probe revealed the presence of dnActRIB transcripts in neurons of the cortex, amygdala, and striatum (data not shown). As expected from the promoter activity (20), the strongest signal was observed in the hippocampal pyramidal and dentate granule cell layers (Fig. 1D). No specific signals were detected with the same probe on sections from control mice (Fig. 1F) or with the sense probe on sections of mice from both genotypes (Fig. 1E and data not shown).

To verify the dominant-negative effect of the transgene in vivo, activin A or vehicle PBS were injected intracerebroventricularly, and the phosphorylation of Smad2 in the ipsilateral hippocampus was monitored by Western blotting 2 h after injection. A basal level of Smad2 phosphorylation was already observed in non-injected or vehicle-injected hippocampi (Fig. 2A), and this signal was predominantly attributed to hippocampal neurons as demonstrated by immunohistochemistry (Fig. 2C). When compared with PBS, activin A reproducibly increased the levels of pSmad2 in the ipsilateral hippocampus of wild-type mice (Fig. 2A), whereas no increase in the levels of pSmad2 were observed in the ipsilateral hippocampus of transgenic mice upon injection of activin A (Fig. 2A). Levels of total Smad2 were not affected by activin A in wild-type and transgenic mice (Fig. 2A). This result was reproduced in three independent experiments with hippocampal lysates from different animals, and the difference between transgenic and wild-type mice was statistically significant (p = 0.012; Fig. 2B). Therefore, the transgene is obviously functional in vivo.

Enhanced Vulnerability to KA Excitotoxicity in dnActRIB Mutant Mice—Histological analysis of the brain of the transgenic animals did not reveal any detectable abnormalities in the non-lesioned hippocampus, and neuronal death was not observed as determined by FJB staining, a marker for degenerating neurons (18) (data not shown). To elucidate the neuroprotective potential of endogenous activin, we performed intracerebroventricular kainate (KA) injections, which cause neuronal loss predominantly in the ipsilateral CA3 region (Fig. 3, A-D, Nissl staining, and E-H, staining with FJB). This injury is accompanied by strong up-regulation of activin betaA expression in wild-type and dnActRIB transgenic mice (Ref. 9 and data not shown). Apoptotic cells were detected by staining with an antibody against cleaved caspase-3 (Fig. 3I). Staining with this antibody correlated with the pattern of FJB staining in the same area (Fig. 3K), although cleaved caspase-3 positive cells comprise only a part of the population of the FJB positive cells. The area of FJB-positive hippocampal cells was measured ipsilaterally and contralaterally in a rectangular area of 1381.5 x 1094.6 µm (Fig. 3, E-H), and every third section through the posterior-anterior extent of the hippocampi was analyzed. Therefore, the data represent an unbiased sampling of FJB-positive cells within the hippocampus. The number of FJB-positive cells in the ipsilateral hippocampi was significantly higher in transgenic (tg) mice of both lines compared with wild-type (wt) littermates (L1-tg, n = 93 sections from 10 mice; L1-wt, n = 100 sections from 10 mice; Pips = 0.0015 (Fig. 3L); L3-tg, n = 60 sections from 6 mice; L3-wt, n = 60 sections from 6 mice, Pips < 0.0001) (Fig. 3M). Furthermore, the extent of cell death in the contralateral hippocampus, the so-called "distant damage" produced by hyperactivity of the commissural fibers connecting the hippocampi was elevated, although the difference was only statistically significant in one mouse line (L3-tg, n = 60 sections from 6 mice; L3-wt, n = 60 sections from 6 mice; Pcon = 0.0004 (Fig. 3M); L1-tg, n = 80 sections from 8 mice; L1-wt, n = 90 sections from 9 mice; Pcon = 0.3697) (Fig. 3L). Since intracerebroventricular KA up-regulates activin betaA mRNA also in the contralateral hippocampus (9), the results suggest that disruption of activin signaling renders hippocampal neurons more vulnerable to both local and distant excitotoxic injury.


Figure 2
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  		FIGURE 2.
dnActRIB is functional in vivo. A, 150 ng of activin A or equal volumes of PBS were injected intracerebroventricularly into wt and tg mice. Ipsilateral hippocampi were isolated 2 h after activin A or PBS injection and analyzed for the presence of pSmad2, total Smad2/3, and beta-actin by Western blotting. B, the levels of pSmad2 in hippocampi of PBS- or activin A-injected wild-type and transgenic mice were densitometrically analyzed. Bars indicate ratio of pSmad2 levels in the ipsilateral hippocampi of activin A-injected versus PBS-injected wild-type mice (2.33 ± 0.38; mean ± S.E., n = 3) and transgenic mice (0.95 ± 0.09; n = 3). Statistical significance was analyzed by Student's t test. *, p < 0.05. C and D, frozen coronal hippocampal sections from untreated hippocampus of wild-type mice were analyzed by immunofluorescence for the presence of pSmad2 (C, red). Staining with the second antibody only is shown in D. Nuclei were counterstained with Hoechst (blue). Note the positive staining for pSmad2 in hippocampal neurons. Bar = 50 µm.

 
The NMDA Receptor-mediated Component of Glutamatergic Neurotransmission Is Reduced in Hippocampi from dnActRIB Mutant Mice—Are neurons from dnActRIB mutant hippocampi more vulnerable to excitotoxic damage, because they respond more sensitively to glutamate? To explore this possibility, we recorded EPSCs from visually identified CA1 pyramidal cells in hippocampal slices. To partially relieve the Mg2+ block of NMDA receptors, extracellular Mg2+ was lowered to 0.3 mM (4.0 mM Ca2+). D-2-amino-5-phosphonovaleric acid (50 µM) and CNQX (10 µM) served to isolate the AMPA and the NMDA receptormediated component of the EPSC, respectively. Whereas the input-output relationship of the overall EPSC was not appreciably altered in mutant hippocampi, the NMDA component was significantly reduced (Fig. 4, A and B; wt, n = 9 slices from 3 mice; L1-dn, n = 9 from 3 mice; L3-dn, n = 8 from 2 mice). The histogram of Fig. 4C shows that the relative contribution of the NMDA current to the overall EPSC (12.9 ± 1.3% in pyramidal cells of normal hippocampi) decreased to 8.1 ± 1.3% and 8.1 ± 0.6% in pyramidal cells of L1 and L3 mutant hippocampi, respectively. By contrast, the time course of NMDA currents did not vary between the different groups (decay time constants: wt, 22.8 ± 2.9 ms, n = 9; L1-dn, 22.3 ± 3.3 ms, n = 9; L3-dn, 23.3 ± 5.3 ms, n = 8, p > 0.9). These data suggest that dnActRIB is unlikely to influence considerably NMDA receptor 2 (NR2) subunit expression, since changes in NR2 subunit composition would be expected to alter the kinetics of NMDA currents (21). A comparison of the expression of NR1, NR2A, and NR2B subunits by means of quantitative real-time RT-PCR lends further support to this notion. Using hippocampal total RNA from wild-type and transgenic mice, we failed to detect a significant change in the expression levels of any of the above subunits (Fig. 5). Furthermore, protein levels of NR2A were not altered as demonstrated by Western blotting of total hippocampal lysates from adult wild-type and transgenic mice (data not shown).

The reduction of the NMDA component was also observed in experiments in which we used bath application of either AMPA (1 µM) or NMDA (10 µM) to mimic the massive rise in extracellular glutamate that is a hallmark of acute excitotoxicity (Fig. 4D). Under these conditions, extrasynaptic glutamate receptors are also activated. In cells of both mutant lines, AMPA responses at more negative potentials were smaller than those in cells from normal hippocampus, but only in L1-dn neurons, this decrease reached statistical significance (Fig. 4E: WT, n = 9 slices from 3 mice; L1-dn, n = 7 slices from 2 mice; L3-dn, n = 8 slices from 3 mice). Since the AMPA receptor-mediated component of the EPSC was not reduced in a similar fashion, it is tempting to speculate that the decreased current responses to AMPA superfusion resulted mainly from an altered expression and/or function of extrasynaptic AMPA receptors. NMDA currents displayed the characteristic increase with membrane depolarization, which was, however, drastically diminished in mutant hippocampi of both lines (Fig. 4F: wt, n = 7 slices from 2 mice; L1-dn, n = 7 slices from 2 mice; L3-dn, n = 8 slices from 3 mice).


Figure 3
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  		FIGURE 3.
dnActRIB-transgenic mice show enhanced vulnerability to KA. Representative Nissl-stained (A-D) or FJB-stained (E-H) frozen coronal brain sections from a transgenic mouse (C, D, G, H) and a wild-type littermate (A, B, E, F) 48 h after KA injection are shown. Bar = 100 µm. Neuronal loss is indicated with arrows. I and K, serial sections from injured hippocampus were immunostained with an antibody against cleaved caspase-3 (I) or FJB (K). Note the similar distribution of cleaved caspase-3- and FJB-positive cells. L and M, quantitative analysis of neuronal cell death in KA-injected wild-type and transgenic mice. The FJB-positive area within the brain region shown in E-H was determined in sections from the ipsilateral and contralateral hippocampus. Bar graphs show mean ± S.E. **, p < 0.01; ***, p < 0.001, n.s. = not significant.

 
Synaptic Plasticity Is Impaired in Hippocampi from dnAc-tRIB Mutant Mice—Since NMDA receptor activation is essential for the induction of LTP in the CA1 region, we wondered whether the decrease of NMDA responses bears significance on the extent of synaptic plasticity in hippocampi of mutant mice. Using a MED64 multielectrode stimulation and recording system, two independent afferent pathways were electrically stimulated to evoke fEPSPs in stratum radiatum of area CA1 (Fig. 6A). During control stimulation, fEPSP slopes did not differ significantly between hippocampi from wild-type and transgenic mice (L1-wt: 111 ± 9 µV/ms, n = 8 versus L1-dn: 86 ± 10 µV/ms, n = 10, p > 0.10; L3-wt: 86 ± 6 µV/ms, n = 18 versus L3-dn 95 ± 5 µV/ms, n = 15, p > 0.23). Also, paired-pulse stimulation (50-ms interstimulus interval) yielded virtually identical ratios of facilitation (L1-wt: 1.51 ± 0.03, n = 16 versus L1-dn: 1.47 ± 0.03, n = 20), p > 0.5; L3-wt: 1.33 ± 0.03, n = 14 versus L3-dn: 1.34 ± 0.05 n = 22, p > 0.5). Following theta burst stimulation, hippocampi from both mutant lines showed significantly impaired LTP compared with controls (Fig. 6B: L1-wt, n = 8 slices from 3 mice; L1-dn, n = 10 slices from 4 mice; Fig. 6C: L3-wt, n = 18 slices from 6 mice; L3-dn, n = 15 slices from 4 mice). Whereas in L1-wt hippocampi, the mean fEPSP slope at 25-30 min post-TBS attained 165 ± 9% of control, L1-dn hippocampi displayed an increase of fEPSP slope to only 135 ± 8% of control (p = 0.033). In recordings from L3 hippocampi, in which extracellular Mg2+ was lowered to 1 mM, the corresponding values were 179 ± 7% for the wt-hippocampi and 155 ± 5% for the dn-hippocampi (p = 0.012). These data indicate that the decrease of the NMDA component observed in whole-cell recordings from mutant hippocampi is sufficient to compromise synaptic plasticity.


Figure 4
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  		FIGURE 4.
NMDA component of glutamatergic transmission is reduced in dnActRIB mutant hippcampi. A, EPSC recordings from CA1 pyramidal cells of wt and L1-tg hippocampi in whole-cell voltage-clamp mode (VH -70 mV, extracellular 0.3 mM Mg2+) show reduced NMDA component after application of the non-NMDA receptor antagonist, CNQX (10 µM), at two stimulus intensities. B, input-output curves for EPSCs from wt and mutant hippocampi. C, histogram of normalized NMDA component in hippocampi from wt and tg mice. EPSC amplitudes before CNQX were about 500 pA. D, inward current responses to AMPA (1 µM) and NMDA (10 µM) superfusion in CA1 neurons of wt and L1-tg mice in the presence of tetrodotoxin (0.5 µM). I-V relationships of AMPA (E) and NMDA responses (F) in hippocampi of wt and tg mice. *, p < 0.05.

 


Figure 5
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  		FIGURE 5.
Expression of NMDA receptor subtypes in the hippocampus. Total RNA was isolated from hippocampi of adult tg and wt littermates and analyzed by quantitative real-time RT-PCR for the mRNA levels of the NMDA receptor subtypes NR1, NR2A, and NR2B. Analysis of beta-actin mRNA levels was used as an internal standard. Bars represent mean ± S.D. from three RT-PCR reactions. The expression level of NR1 was arbitrarily set as 1.

 


Figure 6
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  		FIGURE 6.
LTP is impaired in hippocampi from mutant mice. A, using a multielectrode system, TBS delivered to experimental pathway produced input-specific LTP in area CA1. Insets depict individual fEPSP recordings taken at the like-numbered time points. B and C, compared with hippocampi from wt mice, LTP was significantly reduced in both transgenic lines.

 

   DISCUSSION
TOP
 ABSTRACT
 INTRODUCTION
 EXPERIMENTAL PROCEDURES
 RESULTS
 DISCUSSION
REFERENCES
 
To gain insight into the role of endogenous activin in the normal and injured hippocampus, activin receptor signaling was disrupted in hippocampal neurons. ActRIB was mutated, since it specifically binds to ActRII-activin ligand-receptor complexes (22), and since it is expressed in hippocampal neurons (23).5 Therefore, it can inhibit, in a dominant-negative manner, activin signaling via all activin type I receptors (16). Since expression of the transgene was hardly detectable in cultured hippocampal neurons (data not shown), we confirmed the activity of this mutant and its specificity for activin versus transforming growth factor beta in cultured keratinocytes (24). Most importantly, the dominant-negative effect of the transgene was verified in vivo after intracerebroventricular injection of activin A. In these experiments, activin increased the levels of phosphorylated Smad2 in the hippocampus of wild-type mice but not of transgenic mice. The CaMKII-{alpha} promoter was chosen, because it drives expression of transgenes in forebrain neurons but not in glial cells. Since appreciable activity of this promoter is not observed before the second postnatal week, the transgene is unlikely to interfere with prenatal and early postnatal development (20). Although our study focuses on the role of activin signaling in the hippocampus, it is worth noting that, by using the CaMKII-{alpha} promoter, the transgene is also expressed in other forebrain regions such as cortex, amygdala, and striatum. Consistent with this notion, in situ hybridization indeed revealed the presence of dnActRIB transcripts in these regions. Thus it is well conceivable that the disruption of activin signaling will also bear significance on the normal operation of other brain circuits. Further experiments should help to determine whether activin regulates glutamatergic neurotransmission and excitotoxicity in other forebrain structures in a fashion similar to that reported here from the hippocampus.

Our study demonstrates that activation of activin receptor signaling by endogenous ligands is capable of rescuing hippocampal neurons from excitotoxic cell death. The responsible ligand is most likely activin A, since the betaB chain is not expressed in the normal or lesioned hippocampus (9). A contribution of bone morphogenetic protein 3 (BMP3) to the observed phenotype cannot be fully excluded, since it also binds to activin receptors (25) and since we found a weak expression of BMP3 in normal and injured hippocampus, using real-time RT-PCR (data not shown). However, a contribution of BMP3 to the enhanced vulnerability of the hippocampus of our transgenic mice seems unlikely, since BMP3 is an inhibitor of activin receptor signaling (25) and since activin was shown to be neuroprotective (see above).

Enhanced vulnerability to KA toxicity was observed in both transgenic mouse lines, demonstrating that the effect is not due to integration of the transgene into a host gene, which is important for neuroprotection. Unexpectedly, we found a stronger vulnerability in the transgenic line, which expressed lower levels of dnActRIB. Although the transgene was expressed in forebrain neurons in both mouse lines as determined by in situ hybridization, minor differences in the temporal and spatial expression of the transgene, which are often observed with the CamKII-{alpha} promoter (20), may account for the difference in vulnerability.

Given the selective disruption of activin receptor signaling in neurons, but not in glial cells, neuroprotection by activin receptor ligands is obviously afforded at least in part through a direct action on endangered neurons. Since fibroblast growth factor 2, whose neuroprotective potential is closely linked to the induction of activin A (10), exerts its beneficial action in part by down-regulating NMDA receptors (26), we wondered whether the enhanced vulnerability of dnActRIB mutant mice might be attributable to abnormal NMDA responses. To our surprise, however, we found that under physiological conditions (synaptic stimulation) as well as under pathological conditions (glutamate overflow), the NMDA current was significantly depressed in pyramidal cells of mutant hippocampi, whereas the AMPA current was not or less affected. Thus we discard the hypothesis that the heightened vulnerability of mutant mice to excitotoxic injury results from an overactivity of glutamate receptors. Whereas the neuroprotective mechanisms of activin have thus to await further study, our findings reveal a prominent new function of activin in the normal adult brain that bears particular relevance to synaptic plasticity. The NMDA receptor deficiency that we observed in mature CA1 pyramidal cells of mutant hippocampi should not represent a developmental disturbance, because the transgene is expressed under the control of the CaMKII-{alpha} promoter. We rather propose that activin receptor signaling in adult hippocampus is required to augment NMDA responses, thereby endowing the synapse with its full range of plasticity. Thus genetic or acquired defects of the activin system are likely to impact on synaptic performance in the learning brain as well as on neuronal survival during injury. In the future, it will be important to identify the intracellular signaling pathways, which mediate the effect of activin on forebrain neurons. Since activin was shown to signal via Smad2 in the hippocampus (this study), an involvement of this pathway seems likely. In addition, a role of mitogen-activated kinase signaling should be considered, since activation of ERK1/2 by activin was found to be responsible for the activin-induced expression of tyrosine hydroxylase in mouse E14 primary striatal cell cultures and in a hippocampal neuronal cell line (27).

Our results reveal novel functions of activin at glutamatergic synapses. Thus, in addition to its endocrine functions and its role in inflammation and repair (1, 5), activin seems to be an important modulator of neuronal activity. This finding is likely to be of clinical interest, since application of exogenous activin or stimulation of activin receptors should not only provide protection of endangered neurons but might also emerge as a novel therapeutic approach to improve cognitive impairments.


   FOOTNOTES
 
* This work was supported by the ETH Zurich, the Swiss National Science Foundation (Grant 3100A0-109340/1 (to S. W.)), the University of Kiel, and the Deutsche Forschungsgemeinschaft (SFB 391, TP A9 (to C. A.)). The costs of publication of this article were defrayed in part by the payment of page charges. This article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact. Back

1 Joint senior authors. Back

2 To whom correspondence may be addressed: Institute of Cell Biology, ETH Zurich, Honggerberg, HPM D42, CH-8093 Zurich, Switzerland. E-mail: sabine.werner{at}cell.biol.ethz.ch. 3 To whom correspondence may be addressed: Institute of Physiology, University of Kiel, Olshausenstr. 40, D-24098 Kiel, Germany. E-mail: c.alzheimer{at}physiologie.uni-kiel.de.

4 The abbreviations used are: LTP, long term potentiation; dnActRIB, dominant-negative activin receptor IB; PBS, phosphate-buffered saline; RPA, RNase protection assay; TBS, theta bursts stimuli; AEBSF, 4-(2-aminoethyl)benzenesulfonyl fluoride; RT, reverse transcription; fEPSP, field excitatory postsynaptic potential; FJB, Fluoro-Jade B; KA, kainate; tg, transgenic; wt (or WT), wild-type; EPSC, excitatory postsynaptic current; CNQX, 6-cyano-7-nitroquinoxaline-2,3-dione; NMDA, N-methyl-D-aspartate; NR, NMDA receptor; CaMKII, Ca2+/calmodulin-dependent protein kinase II; BMP, bone morphogenetic protein. Back

5 M. R. Müller and S. Werner, unpublished data. Back


   ACKNOWLEDGMENTS
 
We thank Dr. T. Rülicke for generating the transgenic mice, Dr. P. Bugnon for help with the animal experiments, Dr. C. Bamberger for elp with the characterization of dnActRIB, Drs. C. H. Heldin and P. ten Dijke for the phospho-Smad2 antibody, Dr. I. Mansuy for the CaMKII-{alpha} promoter plasmid, and C. Born-Berclaz for excellent technical assistance.



   REFERENCES
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 ABSTRACT
 INTRODUCTION
 EXPERIMENTAL PROCEDURES
 RESULTS
 DISCUSSION
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