Human Molecular Genetics, 2001, Vol. 10, No. 5 497-505
© 2001 Oxford University Press
Co-regulation of survival of motor neuron (SMN) protein and its interactor SIP1 during development and in spinal muscular atrophy
1Klinische Forschergruppe Neuroregeneration, Department of Neurology, University of Würzburg, Josef-Schneider-Strasse 11, D-97080 Würzburg, Germany, 2Max-Planck Institute of Biochemistry, Am Klopferspitz 18a, D-82152 Martinsried, Germany, 3Institute of Human Genetics, University of Bonn, Wilhelmstrasse 31, D-53111 Bonn, Germany
Received 6 November 2000; Revised and Accepted 11 January 2001.
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ABSTRACT |
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 TOP ABSTRACT INTRODUCTIONRESULTSDISCUSSIONMATERIALS AND METHODSREFERENCES |
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Spinal muscular atrophy (SMA) is a neuromuscular disease characterized by the degeneration of motor neurons in the spinal cord. The disease is caused by mutations of the survival of motor neuron 1 gene (SMN1), resulting in a reduced production of functional SMN protein. A major question unanswered thus far is why reduced amounts of ubiquitously expressed SMN protein specifically cause the degeneration of motor neurons without affecting other somatic cell types. In a first attempt to address this issue we have investigated the Smn interacting protein 1 (Sip1), with an emphasis on its developmental expression and subcellular distribution in spinal motor neurons in relation to Smn. By confocal immunofluorescence studies we provide evidence that a significant amount of Smn does not co-localize with Sip1 in neurites of motor neurons, indicating that Smn may exert motor neuron-specific functions that are not dependent on Sip1. Sip1 is highly expressed in the spinal cord during early development and expression decreases in parallel with Smn during postnatal development. Strikingly, reduced production of Smn as observed in cell lines derived from SMA patients or in a mouse model for SMA coincides with a simultaneous reduction of Sip1. The finding that expression of Sip1 and Smn is tightly co-regulated, together with the unique localization of Smn in neurites, may help in understanding the motor neuron-specific defects observed in SMA patients.
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INTRODUCTION |
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TOPABSTRACTINTRODUCTIONRESULTSDISCUSSIONMATERIALS AND METHODSREFERENCES |
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Spinal muscular atrophy (SMA) is an autosomal recessive disorder characterized by specific degeneration of motor neurons leading to muscle weakness, in particular of proximal voluntary musculature (1). The underlying gene defect on chromosome 5q13 has been identified and characterized. Initial analysis of this chromosomal region revealed two candidate SMA genes, termed survival of motor neuron (SMN) (2) and neuronal apoptosis inhibitor protein gene (NAIP) (3), which are duplicated within this region. One copy of SMN (termed telomeric SMN or SMN1) shows homozygous mutations or deletions in 96% of all SMA patients, whereas the second copy (termed SMN2) is not affected (2,4). Both SMN genes differ in that full-length protein is almost exclusively produced from SMN1. In contrast, the primary transcript of SMN2 undergoes alternative splicing of exon 7, which results in the predominant expression of an unstable and C-terminally truncated SMN protein (2,5,6). At the cellular level, this leads to apoptotic cell death of motor neurons in the anterior horn of the spinal cord and consequently to SMA (1). In contrast to humans, mice harbour only one copy of the Smn gene, which is equivalent to human SMN1. Gene targeting studies revealed that Smn is essential for viability, i.e. mice that are null for the Smn gene die prior to implantation at the blastocyst stage (7). Interestingly, mating of mice with low copy of human SMN2 into the Smn–/– background rescues the early embryonic lethal phenotype but leads to symptoms similar to those found in SMA patients (8,9). A similar phenotype could be observed in mice in which exon 7 of Smn was conditionally deleted in neurons (10). Strikingly, the disease phenotype can be modulated by the copy number of human SMN2 and hence by the relative expression of SMN from this gene (8,9). Specifically, mice that harbour only one or two human SMN2 copies usually develop severe symptoms early in life (corresponding to SMA type I), whereas a less severe phenotype develops in mice with eight SMN2 copies. Thus, SMA is caused by insufficient production of full-length SMN by the SMN2 gene. Moreover, these data also show that the severity of the disease can be modulated by the relative amount of SMN produced from the SMN2 gene locus (8–11).
In the past few years, significant progress has been made in both the identification of SMN-associated proteins and the understanding of their cellular functions. Initial studies revealed that the SMN gene encodes an ubiquitously expressed protein of 294 amino acids, which is found in the cytoplasm and in subnuclear structures of unknown function, termed gemini of coiled bodies (gems) (12). SMN is a Tudor domain protein that is incorporated in large complexes (13). Cellular components that interact with and may hence be functionally connected to SMN include the SMN interacting protein 1 (SIP1) (12,14), the putative DEAD-box helicase dp103/Gemin3 (15,16), the dp103/Gemin3 interacting protein Gemin4/GIP1 (17,18), profilins (19) and spliceosomal U snRNP proteins of the Sm class (12,14,18,20,21). Moreover, SMN transiently interacts with spliceosomal U snRNAs in the cytoplasm of Xenopus laevis oocytes (14).
The association with U snRNAs and Sm proteins suggested a role for SMN in the biogenesis and/or function of spliceosomal U snRNPs (14,20). Indeed, a critical role for SMN and SIP1 in the cytoplasmic assembly of spliceosomal snRNPs U1, U2, U4 and U5 could be deduced from studies in X.laevis oocytes. In this process, the U snRNAs are bound in a step-wise and ordered manner by the Sm hetero-oligomeric protein complexes B/B’.D3, D1.D2 and E.F.G. (22). The U snRNPs are thereafter imported into the nucleus, where they function in pre-mRNA splicing. Antibodies directed against SIP1 and SMN strongly interfere with binding of the Sm proteins on to the U snRNA, indicating that both proteins are essential assembly factors for U snRNPs (14,20). Strikingly, SMA-causing point mutations found in the Tudor domain and the C-terminus of SMN reduced its ability to interact with Sm proteins or to oligomerize (20,21,23). These results raised the possibility that failure of SMN to either oligomerize or to interact with Sm proteins causes, directly or indirectly, the biochemical defects leading to SMA.
Evidence has recently been provided that the nuclear pool of SMN contributes to nuclear pre-mRNA splicing (18,24). Interestingly, in one study inhibition of splicing was observed only after incubation of HeLa nuclear extract with monoclonal antibodies directed against SMN (24). Similarly, pre-incubation of nuclear extract with a deletion mutant of the SMN protein that lacks the first 27 amino acids inhibited splicing by blocking the formation of mature spliceosomes (24). These data indicate that SMN is a component of the pre-mRNA processing machinery that may recycle factors after the splicing reaction. However, it is unclear whether other known SMN interactors such as SIP1, dp103/Gemin3 and Gemin4/GIP1 also contribute to splicing. Nuclear SMN may also have a function related to gene regulation at the transcriptional level. This was concluded from the observation that the papillomavirus nuclear transcription activator E2 interacts with SMN in vitro and in vivo, and that SMN enhances the E2-dependent transcriptional activation of reporter genes (25). Furthermore, the SMN-interacting protein dp103/Gemin3 was originally identified as a factor that interacts with viral and cellular transcription factors (26). Thus, SMN may also regulate gene expression, although the mechanism by which this is achieved is not yet understood in detail.
Despite recent progress in the understanding of the cellular function of SMN, only very little is known about the specific cellular pathology in motor neurons of patients with SMA. In particular, it is unclear why reduced levels of functional SMN protein can lead to specific dysfunction of motor neurons without affecting other cell types and tissues. The proposed cellular role of SMN in U snRNP assembly and splicing suggested that one possible cause for the pathophysiological events in SMA involves defects in pre-mRNA splicing. However, recent findings indicated that the general splicing machinery is unaffected in motor neurons of mice exhibiting symptoms of SMA (27). Hence, the molecular defects leading to motor neuron degeneration in SMA are still unknown.
In this study, we have investigated the cellular localization and expression of Sip1 in relation to Smn in mouse motor neurons. We provide evidence that a significant fraction of Sip1 does not co-localize with Smn in these cells, indicating that Smn may exert motor neuron-specific functions that are not dependent on Sip1. A series of experiments further revealed that the expression level of Smn and Sip1 is tightly co-regulated. Most strikingly, we show that reduced levels of Smn, as observed in cell lines derived from SMA patients or in Smn+/– mice, coincide with a simultaneous reduction of Sip1. These observations may help to understand the pathophysiological events leading to SMA.
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RESULTS |
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Characterization of the murine Sip1 gene
We have isolated a complete cDNA clone of mouse Sip1 by screening a lambda ZAP library with the human cDNA encoding SIP1 (Fig. 1A). The murine Sip1 gene is highly homologous to human SIP1, showing 94% identity at the protein level and 75% at the cDNA level. Using the cDNA as a probe, we isolated a genomic clone that contains the entire coding region of the murine Sip1 gene, including the 5' and 3' flanking regions. The murine Sip1 gene spans a 15 kb region and consists of 10 exons (Fig. 1B). It has recently been reported that the human SIP1 mRNA is alternatively spliced (28). In order to test whether alternative splicing products are also generated from the murine gene, RNA was isolated from mouse 3T3 cells and specific cDNA fragments amplified by primers corresponding to the terminal exon (exon 10) and primers specific for exons 1–9. In all cases, PCR analyses resulted in amplification of only one product of the expected size for a fully spliced Sip1 mRNA (Fig. 1C). Similar results were obtained when splicing of Sip1 was analysed in different tissues (data not shown). This indicates that the mRNA expressed from the murine Sip1 gene is not alternatively spliced and hence encodes only one protein product. This conclusion is further supported by data shown below.
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Sip1 in motor neurons only partially co-localizes with Smn
We next analysed the subcellular localization of Sip1 in motor neurons in relation to Smn. For this purpose, we used a monoclonal antibody directed against mouse Smn and an affinity-purified polyclonal rabbit antiserum raised against recombinant full-length human SIP1 (18). Both antibodies specifically recognize their respective antigen on western blots and in immunoprecipitation experiments (18). To investigate the subcellular distribution of Sip1 in developing motor neurons, the lumbar spinal cord from C57Bl/6 mice was isolated at embryonic day (E)14. Motor neurons were isolated and purified according to procedures established in our laboratory (29) and cultured for 5 days. During this time, the motor neurons grew out axonal and dendritic processes that could easily be recognized in culture (30). Smn and Sip1 in motor neurons were visualized by immunohistochemistry using anti-SIP1 and anti-Smn antibodies, respectively. Initially, laser confocal microscopy was applied to visualize both proteins in the nucleus and the cell body of individual neurons (Fig. 2). Smn and Sip1 could be readily detected in specific nuclear structures previously identified as gems (11) and in the cytoplasm (Fig. 2A and B). Interestingly, an overlay of both images revealed a complete overlap of both proteins in gems but only a partial overlap in the cytoplasm (Fig. 2C). To exclude the possibility that the observed differences in the localization are a specific phenomenon that occurs only in cultured motor neurons, we next investigated the subcellular distribution of both proteins in motor neurons of adult C57Bl/6 mice in situ. Similar to cultured motor neurons, strong Smn and Sip1 immunoreactivity could be observed in the cytoplasm and in nuclear gems of motor neurons (Fig. 2D and E). Importantly, whereas anti-Smn and anti-SIP1 antibodies stained the same gems in the nucleus, immunostaining in the cytoplasm overlapped only partially (Fig. 2D and E). In particular, whereas Sip1 localization was homogenous, Smn could be detected in several distinct dots that were scattered within the cytoplasm. Pre-adsorption of the anti-SIP1 antiserum with recombinant SIP1 strongly reduced the nuclear and cytosolic staining, indicating that the observed signals are highly specific (Fig. 2F). Strikingly, differences in the cytoplasmic distribution were even more obvious when whole motor neurons, including their neurites, were analysed (Fig. 3). Whereas Sip1 was evenly distributed in the cell body and neurites of motor neurons (Fig. 3A–C), Smn appeared highly concentrated in distinct regions of the cell body (Fig. 3D and E) and at branch-points and growth-cones of neurites (Fig. 3F, arrowheads). This differential localization was confirmed by superimposing images stained with anti-Smn and anti-SIP1 antibodies (Fig. 3G–I). Pre-adsorption of the anti-SIP1 antiserum with recombinant SIP1 abolished the signal, indicating the specificity of this serum in immunofluorescence. Thus, our data indicate that a significant fraction of cytosolic Smn does not co-localize with, and may hence exist unbound to Sip1 in neurites of motor neurons. This indicates that Smn and Sip1 serve functions in these cells that are not strictly dependent on each other.
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Differential expression of Sip1 in tissues of adult mice
In order to investigate Sip1 expression in different tissues, protein extracts from liver, lung, heart, kidney, spleen, muscle, spinal cord and brain of adult C57Bl/6 mice were isolated and separated by denaturing sodium dodecyl sulphate–polyacrylamide gel electrophoresis (SDS–PAGE). The proteins were transferred to a nitro-cellulose membrane and SIP1 was visualized by western blotting using the same polyclonal anti-SIP1 antibody as above. Sip1 was detected in all tissues as a single band, further confirming that transcripts from the Sip1 gene are not alternatively spliced (Fig. 4). The additional 35 kDa band visible in muscle tissue is a non-specific cross-reactivity of the antiserum. Interestingly, the expression level of Sip1 protein varies considerably between different tissues. As shown in Figure 4 (top panel), high levels of Sip1 protein are present in liver, kidney, spleen and brain, whereas significantly lower amounts were detected in lung, heart, muscle and spinal cord. In this respect, the expression of Sip1 and Smn in different tissues of adult mice is remarkably similar to Smn (compare top and middle panels). All blots were stripped and reprobed with an anti-actin antibody to ensure that equal amounts of proteins were loaded on to the gel (bottom panel).
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Downregulation of Sip1 in motor neurons of Smn+/– mice and in cell lines derived from SMA patients
The expression of similar amounts of Smn and Sip1 in different tissues suggested that both proteins are co-regulated at the level of expression or turnover. To test this possibility, we compared the amounts of Smn and Sip1 in the spinal cord during development (Fig. 5). Sip1 expression was highest in the developing spinal cord between E14 and birth (E19) and expression decreased during postnatal development [postnatal day (P)15, 5 weeks, 1 year]. Interestingly, the decrease of Sip1 goes in parallel with Smn, which also shows the highest relative expression during embryonic development but lower levels during later postnatal phases (Fig. 5).
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The above results implied that cells with reduced levels of SMN, such as cells from patients suffering from SMA, also express lower levels of SIP1. The availability of cell lines derived from SMA patients and tissues from Smn+/– mice allowed us to test this possibility directly. Initially, we determined the amount of Smn and Sip1 in the spinal cord of 12-month-old heterozygous mice lacking one Smn allele (Smn+/– mice) (Fig. 6). In comparison with their normal littermates, these mice have Smn protein levels that are reduced by
50%, consistent with the lack of one out of two Smn alleles (27). Strikingly, the spinal cord of these mice also contains 50% less Sip1 protein compared with the spinal cord of wild-type mice (Fig. 6A and C). Importantly, the reduction of Sip1 protein in Smn+/– mice is not only detectable in spinal cord but also in other tissues such as muscle and brain (Fig. 6B). Thus, the artificial reduction of Smn in Smn+/– mice induces a general and simultaneous downregulation of Sip1 by roughly the same factor.
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Finally, we investigated the expression of SMN and SIP1 in cells derived from patients suffering from SMA (Fig. 7A and B). Proteins were extracted from Epstein–Barr virus (EBV)-transformed lymphoblastoid cell lines derived from either type I SMA patients (n = 3) or unaffected controls (n = 3) and analysed by western blotting with anti-Smn and anti-SIP1 antibodies. As expected, these cells express less SMN than those from non-affected individuals due to deletions in the SMN1 locus (Fig. 7A, middle panel). Importantly, in parallel with a reduction of SMN, we also detected less SIP1 protein in these cells (Fig. 7A, top panel; for quantification see also Fig. 7B). In conclusion, these data strongly suggest that the expression of SMN and SIP1 is tightly co-regulated in patients suffering from SMA.
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DISCUSSION |
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TOPABSTRACTINTRODUCTIONRESULTSDISCUSSIONMATERIALS AND METHODSREFERENCES |
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Although genetic studies provided strong evidence that mutations in the SMN1 gene locus are the primary cause of SMA, a major question unanswered thus far is why reduced amounts of ubiquitously expressed SMN protein specifically cause the degeneration of motor neurons without affecting other somatic cell types. One explanation for this paradox is that SMN, in addition to its reported functions, exhibits motor neuron-specific activities that are unimportant for the normal functioning of other cells. These activities may be brought about by SMN itself or by other factors that interact with SMN in a cell type-specific manner. In a first attempt to test this hypothesis, we have focused our studies on the expression and intracellular localization of SIP1 in relation to SMN.
Several lines of evidence presented here indicate that the expression levels of Smn and Sip1 are co-regulated. First, the expression pattern in different tissues is remarkably similar, i.e. those tissues that express large amounts of Smn (liver, kidney, spleen and brain) also show the highest level of Sip1. Conversely, tissues with only low levels of Smn (heart, muscle and spinal cord) also show low expression of Sip1. Second, Smn and Sip1 are downregulated in the developing spinal cord with similar kinetics. Specifically, expression of both proteins is high in early embryonic development and gradually decreases as development progresses. Third, the non-physiological reduction of SMN by 50% in Smn+/– heterozygous mice results in a simultaneous decline of Sip1. Fourth, and most importantly, SMA type I patients not only exhibit reduced levels of SMN, but also show low expression levels of SIP1 in comparison with unaffected individuals. Thus, we provide evidence in different model systems that the reduced expression of Smn coincides with the simultaneous reduction of Sip1. Whether this co-regulation of Smn and Sip1 occurs at the level of protein stability, transcription and/or translation is currently under investigation. Given that SMN is found in macromolecular complexes that contain in addition to SIP1 a variety of other proteins (12,15–19), it is a possibility that the expression of other SMN-interacting proteins is also subject to similar regulatory events. These results are likely to be of importance for understanding the molecular events leading to SMA as they suggest that apart from SMN, the expression level and hence the activity of other SMN associated proteins may be altered in SMA patients. If co-regulation of SMN interacting factors in SMA patients is a general feature and also affects putative factors that interact with SMN in a cell type-specific manner, this could give a first clue as to why the disease phenotype manifests itself only in motor neurons. Most known SMN interactors such as SIP1, Gemin3/dp103 and Gemin4/GIP appear to be ubiquitously expressed and hence are unlikely to mediate cell-specific functions. On the other hand, profilin II, which has recently been shown to interact with SMN, is predominantly expressed in motor neurons of anterior horns (19). It will therefore be of major interest to investigate the localization of profilin II in neurites and its expression level in response to reduced production of SMN.
In somatic cells, SMN forms a tight complex with SIP1 and both proteins completely co-localize in the nucleus and the cytoplasm (11). Consequently, it has been assumed that most or possibly all SMN is bound to SIP1 in vivo. Strikingly, however, this appears not to be the case for Smn in motor neurons as in these cells a significant fraction does not co-localize and is hence likely not to be bound to Sip1. These studies give a first indication for motor neuron-specific function(s) of Smn and may provide a link between reduced levels of SMN and the pathophysiological events leading to SMA. Interestingly, Smn that is unbound to Sip1 is found predominantly in axons and dendrites, i.e. at the very periphery of motor neurons. In contrast, most Smn found in the cell body is likely to be in a complex with Sip1, since both proteins co-localize in this region [this study, see also Battaglia et al. (31)]. For several reasons, it is unlikely that SMN that localizes in axons or in dendrites plays a role in the biogenesis of U snRNPs. First, it has been shown that U snRNP assembly involves a complex containing both SMN and SIP1 (14,20). Regions within axons and dendrites that lack stoichiometric amounts of Sip1 are therefore unlikely to be sites of assembly. Second, earlier studies indicated that U snRNAs are only transiently exported from the nucleus to assemble in the cytoplasm (32). Hence, it is believed that the biogenesis of U snRNPs primarily occurs in the cell body and not in structures that are at a considerable distance, such as dendrites or axons. Together, these data raise the possibility that SMN fulfils a motor neuron-specific function in axons and dendrites that is not dependent on SIP1. The function of Smn in these neuronal structures is currently unknown. Likewise it is an open question whether profilin II or other as yet unidentified proteins are involved in this function. To this end, the biochemical purification of protein complexes containing SMN from motor neurons should reveal tissue-specific factors that are not found in complexes of non-neural cells. The availability of elaborate purification protocols for the isolation of SMN complexes, as reported by two groups (16,18), will further help to answer the question of why motor neurons are the primary target in SMA.
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MATERIALS AND METHODS |
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TOPABSTRACTINTRODUCTIONRESULTSDISCUSSIONMATERIALS AND METHODSREFERENCES |
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Western blot analysis
Heart, liver, lung, kidney, spleen, muscle, brain and spinal cord were dissected from Smn+/+ and Smn+/– mice (derived from the same litters). The tissues were homogenized and the extracts dissolved in RIPA buffer (50 mM Tris pH 7.5, 150 mM NaCl, 1% Nonidet P-40, 0.5% sodium deoxycholate, 0.1% SDS). After centrifugation, the protein concentration of the supernatants was determined using the Bio-Rad Protein Assay kit (Bio-Rad) according to the manufacturer’s instructions. EBV-transformed lymphoblastoid cell lines from SMA patients and control persons were cultured according to standard protocols and used to prepare protein extracts. Each protein extract was mixed with the same volume of sample buffer (125 mM Tris pH 6.8, 4% SDS, 10% ß-mercaptoethanol, 20% glycerol, 0.004% bromophenol blue). The samples were boiled for 2 min and stored at –20°C, and then electrophoresed on a 10% polyacrylamide gel, transferred to Protean membrane (Schleicher & Schuell) with the Biometra Fastblot system B24 at 7 V/cm constant voltage for 1 h in 25 mM Tris pH 8.3, 150 mM glycine and 10% methanol. Unspecific binding sites of the Protean membrane were blocked for 30 min with 5% instant milk in TBS containing 0.2% Tween (TBS-T). Primary antibodies including the monoclonal anti-mouse Smn IgG1 (250 µg/ml, Dianova), the polyclonal rabbit antiserum directed against human anti-SIP1 (18) and anti-actin (1mg/ml) (Roche) were diluted 1:1.000 in 5% instant milk in TBS-T. The anti-Smn antibody reacts with full-length mouse Smn protein but not with truncated Smn lacking the N-terminus including the Sip1 interaction domain (S. Jablonka and M. Sendtner, unpublished data). The antibodies were incubated with the membrane for 2 h at room temperature. The membrane was washed three times for 15 min at room temperature with TBS-T. Goat anti-mouse and goat anti-rabbit HRP conjugated antibodies (Roche) were used as secondary antibodies at a dilution of 1:10.000 with 5% instant milk in TBS-T. The membrane was incubated for 1 h at room temperature and subsequently washed three times. Then, the Smn immunoreactive bands were visualized using ECL chemiluminescent reagent (Amersham) according to the manufacturer’s instructions. The blots were exposed to Fuji medical X-ray film (Super RX) for the detection of the chemiluminescent emissions. Each experiment was repeated at least twice, Smn immunoreactive bands scanned and the intensity quantified, using the Aida Software 2.0x (Raytest). Stripping and reprobing of the blots was performed according to the manufacturer’s protocols.
Immunodetection of Smn and Sip1 in spinal motor neurons from spinal cord sections
The lumbar spinal cord from a 12-month-old C57Bl/6 mouse was dissected and the L4 segment identified and frozen in Tissuetec (Sakura). Frozen sections (10 µm) were prepared, blocked with TBS containing 10% BSA for 20 min at room temperature and incubated with a monoclonal antibody against mouse Smn (Dianova) at 1 µg/ml and polyclonal antibodies against human SIP1 (1:1000). The samples were then washed three times with TBS, incubated with secondary antibodies [anti-mouse IgG fluorescein conjugate (Roche Molecular Biochemicals) and CyTM3-conjugated goat anti-rabbit (IgG, Dianova)], washed again three times with TBS, embedded with DABCO (Merck) and covered with glass slides. Smn and Sip1 immunoreactivity was visualized with a Leica confocal microscope. The settings for pinhole and voltage were identical for the analysis of all sections. This experiment was repeated four times; data shown are from one representative experiment.
Immunodetection of Smn and Sip1 in mouse embryonic motor neurons
Pregnant mice (14 days after conception) were killed by cervical dislocation and embryos were dissected from the uterus and decapitated prior to further handling. Cultures of spinal motor neurons from E14 mice after enrichment for motor neurons were prepared by a metrizamide cushion centrifugation technique (33). The ventrolateral parts of the lumbar spinal cord were dissected and transferred to HBSS containing 10 µM ß-mercaptoethanol. After treatment with trypsin (0.05%, 10 min), tissues were triturated and the cell suspension passed through a nylon mesh (100 µm pore size). The cells were overlaid on 10% metrizamide in HBSS. The metrizamide cushion was centrifuged for 20 min at 400 g and cells from the interphase were removed and transferred to culture medium without apotransferrin. Cells were plated at a density of 2000 cells/cm2 in 4-well culture dishes (Greiner) precoated with poly-ornithine and laminin as described by Arakawa et al. (34). Cells were grown in neurobasal medium (Gibco) with serum and 500 µM glutamax and 50 µg/ml apotransferrin at 37°C in a 5% CO2 atmosphere. Fifty percent of the medium was replaced at day 1 and then every second day.
Motor neurons kept in culture for 5 days were fixed with methanol/aceton (1:1) and incubated with a mouse monoclonal antibody against Smn (Dianova) at 1 µg/ml and a polyclonal antiserum against SIP1 (1:1.000). The samples were washed three times with TBS, incubated with secondary antibodies [CyTM3-conjugated goat anti mouse IgG, (Dianova) and anti-rabbit IgG fluorescein conjugate (Roche Molecular Biochemicals)], washed again three times with TBS and embedded with Mowiol. Immunoreactivity was visualized with a Leica confocal microscope. The settings for pinhole and voltage were identical for the analysis of all motor neurons.
Cloning of genomic and cDNA sequences of SIP1
The genomic sequence of murine Sip1 was isolated from a BAC library (Genome Systems) of a mouse RW4-cell line (mouse strain 129SVJ). Briefly, the library was screened with the hSIP1 cDNA. Positive clones were subcloned in pBS-KS and sequenced. Oligonucleotides encompassing the 5'- and 3'-end of the coding sequence of murine Sip1 gene were used to isolate a full-length Sip1 cDNA from a cDNA library. The authentic 5'-end was further confirmed by 5' race via RT–PCR with RNA from mouse 3T3 cells. All enzymatic reactions were carried out with standard protocols as provided by the manufacturers/distributors. For detection of alternatively spliced Sip1 products, a cDNA template was synthesized via RT–PCR using an oligo-dT23 primer for the reverse transcription and mSip1 specific primers specific for exon-1 (matching nucleotides 59–94) and reverse primers (matching nucleotides 1126–1091) of exon 10. In a second round, this template was used for a nested PCR with forward primers specific for either exon 1 (nucleotides 196–231), 2 (nucleotides 261–296), 3 (nucleotides 340–375), 4 (nucleotides 427–462), 5 (nucleotides 541–576), 6 (nucleotides 592–626), 7 (nucleotides 645–680) or 8 (nucleotides 744–778) and a reverse primer matching exon 10 (nucleotides 1050–1021). PCR products were detected on conventional 1.2% agarose gels containing ethidium bromide.
RNA used to perform RT–PCR studies was isolated by lysing L1 mouse 3T3 cells (1–5 x 106 cells) using 500 µl of a solution containing 3 M guanidinium-thiocyanate, 10 mM ß-mercaptoethanol and 10 mM EDTA, followed by an acidic phenol–chloroform extraction. The RNA was subsequently ethanol-precipitated and resuspended in water. Sequence homology database screens and homology fits were performed using NCBI BLAST or the Wisconsin Package versions 9.1 and 10.0 of the Genetic Computer Group.
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ACKNOWLEDGEMENTS |
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We are indebted to J. Kara, F. Schoenen, H. Raschke and G. Sowa for excellent technical assistance. This work was supported by grants from the Deutsche Forschungsgemeinschaft to M.S. (SFB 581/TP B1), U.F. (DFG Fi-573/2-1) and B.W. (SFB400/TP A6) and families of SMA. U.F. receives support from the Max-Planck Society.
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FOOTNOTES |
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+ These authors contributed equally to this work.
§ To whom correspondence should be addressed. Tel: +49 89 8578 2475; Fax: +49 89 8578 3965; Email: ufischer@biochem.mpg.de
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REFERENCES |
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