Human Molecular Genetics, 2001, Vol. 10, No. 26 3017-3024
© 2001 Oxford University Press
SALL1, the gene mutated in Townes–Brocks syndrome, encodes a transcriptional repressor which interacts with TRF1/PIN2 and localizes to pericentromeric heterochromatin
Institute of Human Genetics, University of Göttingen, Heinrich-Düker-Weg 12, 37073 Göttingen, Germany and 1Institute of Anthropology and Human Genetics, University of Munich, Richard-Wagnerstraße 10/I, 80333 München, Germany
Received August 20, 2001; Revised and Accepted October 26, 2001.
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ABSTRACT |
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 TOP ABSTRACT INTRODUCTIONRESULTSDISCUSSIONMATERIALS AND METHODSREFERENCES |
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The Townes–Brocks syndrome (TBS) is an autosomal dominantly inherited malformation syndrome presenting as an association of imperforate anus, triphalangeal and supernumerary thumbs, malformed ears and sensorineural hearing loss. Mutations in SALL1, a gene mapping to 16q12.1, were identified as a cause for TBS. To elucidate how SALL1 mutations lead to TBS, we have performed a series of functional studies with the SALL1 protein. Using epifluorescence and confocal microscopy it could be shown that a GFP–SALL1 fusion protein localizes to chromocenters and smaller heterochromatin foci in transiently transfected NIH-3T3 cells. Chromocenters consist of clustered pericentromeric heterochromatin and contain telomere sequences. Indirect immunofluorescence revealed a partial colocalization of GFP–SALL1 with M31, the mouse homolog of the Drosophila heterochromatic protein HP1. It was further demonstrated that SALL1 acts as a strong transcriptional repressor in mammalian cells. Transcriptional repression could not be relieved by the addition of the histone deacetylase inhibitor Trichostatin-A. In a yeast two-hybrid screen we identified PIN2, an isoform of telomere-repeat-binding factor 1 (TRF1), as an interaction partner of SALL1, and showed that the N-terminus of SALL1 is not necessary for the interaction with PIN2/TRF1. The interaction was confirmed in vitro in a GST-pulldown assay. The association of the developmental regulator SALL1 with heterochromatin is striking and unexpected. Our results propose an involvement of SALL1 in the regulation of higher order chromatin structures and indicate that the protein might be a component of a distinct heterochromatin-dependent silencing process. We have also provided new evidence that there is a close functional link between the centromeric and telomeric heterochromatin domains not only in Drosophila and yeast, but also in mammalian cells.
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INTRODUCTION |
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TOPABSTRACTINTRODUCTIONRESULTSDISCUSSIONMATERIALS AND METHODSREFERENCES |
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Townes–Brocks syndrome (TBS; OMIM 107480) is an autosomal dominantly inherited malformation syndrome. Mutations in SALL1, a gene mapping to 16q12.1, were identified as the cause for TBS (1). The clinical presentation of TBS is highly variable within and between affected families (2). Characteristic features of TBS are anorectal abnormalities (imperforate anus, anal stenosis), abnormalities of the hands (preaxial polydactyly, triphalangeal thumbs), abnormalities of the feet (syndactyly, pes planus, fused metatarsals), deformities of the outer ear (‘lop ears’, microtia), preauricular tags and hearing loss, which can be sensorineural, conductive or mixed. Renal malformations have also been reported from several cases and can lead to renal failure in TBS patients. Cardiac malformations as well as mental retardation are rarely reported. In addition, some families are known in which affected subjects show features typical for both TBS and the Goldenhar syndrome/oculo-auriculo-vertebral spectrum (OMIM 164210). Penetrance seems to be complete in TBS (2).
SALL1 has homologies to the essential developmental regulator gene spalt (sal) of Drosophila (3). In Drosophila, sal is required for the specification of posterior head and anterior tail segment identity (4) as well as for larval tracheal system development (5) and adult wing development (6). In the wing imaginal discs, sal is activated in response to hedgehog signalling mediated by the TGFß-like protein DPP (7–9).
Prior to the cloning of SALL1, sal-related genes had been isolated from mouse (Msal, now Sall3) (10) and from Xenopus laevis (Xsal-1) (11). Both genes are expressed in the developing limbs, heart, kidney, inner ear and central nervous system, i.e. in tissues/organs affected in TBS. Human SALL1 is very similar in structure to both Msal and Xsal-1, and shows a similar expression pattern in adult tissues (12). The observation that the sal gene in the fish Medaka is activated in response to Sonic hedgehog suggests that not only the SAL-like protein structure was conserved in evolution but that SAL-like proteins might also act as essential developmental regulators in vertebrates (13).
The ORF of SALL1 encodes a protein of 1325 amino acids with four characteristically arranged SAL-like C2H2 double zinc finger domains (Fig. 1). A single zinc finger is attached to the second double zinc finger domain (12). All known vertebrate SAL-like proteins contain an additional C2HC zinc finger close to the N-terminus (11–14). The majority of the SALL1 mutations detected in TBS patients to date result in preterminal stop codons (12). It is very likely that TBS is caused by a haploinsufficiency of SALL1. Among typical TBS patients, we reached a detection rate of SALL1 mutations of 64.3% (15). However, there are a number of families with typical TBS-like phenotypes in which SALL1 mutations were not found.
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To elucidate how SALL1 mutations lead to TBS, we performed a series of functional studies with the SALL1 protein. First, we determined the intracellular localization of a GFP–SALL1 fusion protein in NIH-3T3 cells by epifluorescence and confocal microscopy. Secondly, we tested SALL1 for its ability to repress or activate gene expression in transient transfection assays. Thirdly, we performed a yeast two-hybrid-screen to identify proteins interacting with SALL1.
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RESULTS |
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SALL1 colocalizes with heterochromatin
NIH-3T3 cells (mouse fibroblasts) were transfected transiently with pEGFP–SALL1 to determine the intracellular localization of the fusion protein in interphase cells. By epifluorescence microscopy, we found an exclusive nuclear localization of GFP–SALL1 in almost all cells. However, in some cells GFP–SALL1 was also present in the cytoplasma (data not shown). The cytoplasmatic fraction often looked like inclusion bodies and possibly represented membrane-bound vesicles that were formed due to the relative overexpression in certain cells. In all transfected cells (even in those few cells with a cytoplasmatic GFP–SALL1 fraction) GFP–SALL1 was distributed in the nucleus as distinct aggregates (Fig. 2A). These aggregates strikingly corresponded to the 4',6-diamidin-2-phenylindol (DAPI)-bright regions of the nucleus (Fig. 2B). The DAPI-bright regions in the nuclei of NIH-3T3 cells represent the chromocenters and consist of clustered pericentromeric heterochromatin.
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Confocal microscopy was used in order to analyse the pattern of the GFP–SALL1 distribution at a higher resolution. Heterochromatin (and hence chromocenters) were stained with TO-PRO®-3. This technique revealed distinct distribution patterns of GFP–SALL1: in the nucleus depicted in Figure 2C and D GFP–SALL1 shows the previously described colocalization with the chromocenters, which were located at the nuclear membrane or at the periphery of the nucleoli. In the nucleus shown in Figure 2E and F GFP–SALL1 not only colocalizes with the chromocenters, but is also distributed in smaller foci which seem to cover the inner surface of the nuclear membrane and the outer surface of the nucleoli. A tangential section of such a cell (Fig. 2G) reveals that SALL1 covers the inner surface of the nuclear membrane in a mesh-like fashion. The nuclei depicted in Figure 2A–F show a homogenous distribution of GFP–SALL1 within the chromocenters. However, in other nuclei a pattern was observed which looked like coating of the outer surface of the chromocenters (Fig. 2H–J). In these cells, the coating did not cover the complete surface of the chromocenters, but contained openings.
SALL1 partially colocalizes with M31/HP1
Since the mammalian homologs of Drosophila heterochromatin protein 1 (HP1) are known to localize to constitutive heterochromatin, we used indirect immunofluorescence with a monoclonal antibody (raised against M31, the murine HP1 homolog) to stain the pericentromeric heterochromatin. Epifluorescence microscopy (Fig. 2K–M) and confocal microscopy (Fig. 2N–P) revealed that GFP–SALL1 and M31/HP1 partially colocalize at the chromocenters and showed a M31/HP1 distribution in the nucleus as described by Minc et al. (16) and Wreggett et al. (17). Comparing the distribution patterns of the two proteins in detail showed that most subregions of the chromocenters are either exclusively covered by GPP–SALL1 or by M31/HP1.
SALL1 acts as a transcriptional repressor in mammalian cells
Since one of the hallmarks of heterochromatin is that it constitutes a transcriptionally repressive environment, we next wished to examine whether SALL1 acts as a transcriptional repressor. To this end, SALL1 was tested for its transcriptional repression properties in a reporter gene assay. A series of transient transfections of NIH-3T3 cells was performed with a full-length SALL1 fusion protein (Fig. 1) with the GAL4-DNA-binding domain (GAL4-DB). As a reporter plasmid we used pGAL45tkLUK, which contains the luciferase gene under the control of a thymidine kinase promoter with a GAL4 binding site. The results of these assays are shown in Figure 3. GAL4-DB–SALL1 very strongly represses the activity of this promoter. Luciferase expression was repressed 
20-fold (Fig. 3, lane 3). We next tested if transcription repression of SALL1 is relieved by the addition of the histone deacetylase inhibitor Trichostatin-A (TSA). As a control the central domain (amino acids 218–345) of the ets transcription factor ETV6 (Fig. 3, lane 5) was used. Transcription repression of the ETV6 central domain is known to be mediated through histone deacetylases (18) (S.K.Bohlander, J.Putnik, S.Bartels and M.Kickstein, manuscript in preparation). The results, shown in Figure 3, lane 4, clearly indicate that TSA had no effect on the transcription repression of SALL1. However, the repression by the central domain is relieved in the presence of TSA (Fig. 3, lane 6).
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SALL1 interacts with PIN2 in yeast
The yeast two-hybrid system was applied to identify proteins that interact with SALL1 using pGBT9–SALL1
2 (encoding amino acids 87–1058 of SALL1; Fig. 1) as a bait. Among 2 x 106 transformants we identified one clone that was positive for the expression of the selection markers (ß-galactosidase and histidine). Subsequent sequence analysis showed that the clone had a 792 bp insert containing an open reading frame (ORF) of 264 amino acids. A database search revealed 100% homology with the human PIN2 protein (amino acids 67–331; Fig. 4), an isoform of telomere repeat binding factor 1 (TRF1).
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To determine the domains of SALL1 necessary for the interaction with PIN2, pGAD-GH–
PIN2 was cotransformed with pGBT9–SALL1, –SALL12, –SALL13 and –SALL1 (Fig. 1) into yeast strain CG1945 and assayed for ß-galactosidase and histidine activity. Full length SALL1, SALL12 and SALL13, but not SALL11 interacted with PIN2 (Fig. 5; data for complete SALL1 not shown). This indicates that the N-terminus of SALL1 is not necessary for interaction with PIN2.
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SALL1 interacts with PIN2 in vitro
To confirm that SALL1 and PIN2 interact physically, we performed a GST-pulldown assay. The fusion protein GST–
PIN2 (amino acids 67–331 of complete PIN2; Fig. 4) was expressed, purified and incubated with in vitro translated 35S-Met-labeled SALL12. After extensive washing, the proteins bound to glutathione sepharose beads were separated by SDS–PAGE and detected by autoradiography. The result of the assay is shown in Figure 6: GST–PIN2 (lane 2), but not GST alone (lane 3) is able to efficiently retain SALL12.
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DISCUSSION |
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Using epifluorescence microscopy we have shown that in interphase NIH-3T3 mouse fibroblasts the GFP–SALL1 fusion protein is concentrated in DAPI-bright regions of the nucleus, which are known to represent chromocenters. The chromocenters of mouse nuclei consist of pericentromeric heterochromatin (19) and contain some telomeres, since mouse chromosomes are acro- or telocentric (20–22). Confocal microscopy with TO-PRO®-3 heterochromatin staining confirmed the observed SALL1 localization and revealed that GFP–SALL1 exhibits different distribution patterns within the nuclei: in some cells the protein was not only localized at the chromocenters, but was also distributed in smaller foci which seemed to cover the inner surface of the nuclear membrane and the outer surface of the nucleoli. These differences might reflect cell-cycle dependent changes in heterochromatin organization and hence GFP–SALL1 localization. Interestingly, we never observed a GFP–SALL1-transfected cell in mitosis or arrested in metaphase after colchicin treatment. Therefore, overexpression of SALL1 might either be toxic or lead to cell-cycle arrest. Further experiments are needed to interpret this observation and to examine the localization of native SALL1 at different stages of the cell cycle. Nevertheless, at least three independent lines of evidence strongly argue against the possibility that the observed association of SALL1 with heterochromatin is an artefact of the overexpression of the fusion protein. First, GFP alone shows a diffuse cytoplasmic and nuclear localization without any accumulation at DAPI-bright regions in the nucleus (data not shown). Secondly, the localization of GFP–SALL1 is highly specific; the protein is (in the nucleus) exclusively found at heterochromatic sites. Thirdly, we identified a SALL1 interaction partner (PIN2/TRF1) that binds telomeric heterochromatin and also localizes to chromocenters (22,23).
Taking into account that mutations of SALL1 lead to a well defined and specific phenotype, the accumulation of SALL1 at sites of highly repetitive DNA is striking and unexpected. Our results suggest that SALL1 is not a ‘classical’ transcription factor regulating the expression of a few target genes, but that the protein is involved in the regulation of higher order chromatin structures such as pericentromeric heterochromatin. There are only two other examples of human malformation syndromes that are caused by mutations in a gene coding for a heterochromatin-associated protein. (i) The ATR-X syndrome, a severe form of syndromal mental retardation characterized by the presence of 
-thalassemia with urogenital abnormalities and facial dysmorphism, is caused by mutations of the X-chromosomal gene hATRX. The ATRX protein is a putative transcriptional regulator which is localized at the pericentromeric heterochromatin and at the short arms of acrocentric chromosomes (24). (ii) The ICF syndrome (an acronym for immunodeficiency, centromeric instability and facial anomalies), an autosomal recessive disorder, is caused by mutations in the DNA methyltransferase DNMT3B and involves extensive loss of methylation from pericentromeric regions (25). Just recently it was shown that DNMT3B is a methylation-independent transcriptional repressor and colocalizes with HP1 to pericentromeric heterochromatin regions in murine embryonic stem cells (26).
We have shown that SALL1 is a strong transcriptional repressor when it is tethered to a promoter. This effect is apparently not mediated by histone-deacetylases. The exact repression mechanism remains to be determined. Since the SALL1 mutations detected in Townes–Brocks syndrome are believed to result in haploinsufficiency, the Townes–Brocks phenotype might be caused by the up-regulation of SALL1 target genes. The fact that SALL1 is located at heterochromatin regions suggests that the protein is involved in the establishment or stabilization of a transcriptionally repressive environment. Changes in chromatin organization and the directed formation of heterochromatin-like complexes are thought to be important for regulating gene expression during development (27). For example, Ikaros, another zinc finger protein that binds centromeric heterochromatin, is required for normal T, B and natural killer cell development and is thought to repress genes by selectively recruiting them to centromeric heterochromatin (28). Taking into account the fact that SALL1 exhibits a spatiotemporally restricted expression profile, our findings indicate that SALL1 might be a component of such a distinct heterochromatin-dependent silencing process.
A similar clustering at chromocenters as observed for SALL1 has been reported for M31, the murine homolog of the Drosophila heterochromatic protein HP1. Therefore, we visualized HP1/M31 in GFP–SALL1 transfected cells by indirect immunofluorescence. As would be expected from the published data on the intracellular localization of HP1/M31 (16,17), the signals detected from both proteins partially overlapped at DAPI-bright regions of the nucleus. In addition, HP1/M31 staining revealed smaller spots that were not overlapping with GFP–SALL1. These signals most likely represent unspecific background, as they were also detectable in the cytoplasma (data not shown). Interestingly, most subregions of the chromocenters were either covered exclusivly by SALL1 or by HP1. We did not perform any experiments to test whether the two proteins directly interact with each other. HP1 was first described in Drosophila (29) as a heterochromatin-associated protein with dosage-dependent effects on heterochromatin-induced gene silencing known as position effect variegation (30). In Drosophila, HP1 is also associated with telomeres and is thought to be involved in preventing telomere fusions (31). In Schizosaccharomyces pombe, the swi6 gene encodes a HP1-like chromodomain protein that localizes to heterochromatin domains, including the centromeres and telomeres, and is involved in silencing at these loci (32). A number of interactions with other proteins have been reported for the mammalian homologs of HP1, among which the interaction of HP1 with human Ku70 is especially intriguing (33). Ku, which is a heterodimer of 70 and 80 kDa subunits, is involved in DNA repair and the maintenance of telomeres (34–36). Ku70 also interacts with TRF2, a mammalian telomere-binding protein (37). It has been suggested that HP1 is a mammalian counterpart of the yeast Sir4 protein and represents a telomere protein of the silencing group (33).
There seem to be some striking parallels between HP1 and SALL1 not only concerning their intracellular localization, but also regarding functional aspects, since our yeast two-hybrid screen identified a SALL1 interaction partner which also acts at telomeres. PIN2 is a splice variant of the telomere-repeat-binding protein TRF1, carrying an internal 20 amino acid deletion (23). TRF1/PIN2 is involved in regulating telomere length. Long-term overexpression of TRF1 in a telomerase-positive tumor cell line results in progressive telomere shortening, whereas inhibition of TRF1 induces telomere elongation (38). PIN2 is more abundant in cells than TRF1, but functional differences between PIN2 and TRF1 have not been reported. So far, TRF1 protein interactions with Tankyrase, TIN2 and NBSI have been reported. Tankyrase, a protein with homology to ankyrins and the catalytic domain of poly(ADP-ribose) polymerase, colocalizes with telomeres and can ribosylate both itself and TRF1 in vitro (39). Ribosylation of TRF1 inhibits its binding to telomeric DNA. TIN2 is thought to mediate TRF1 function and negatively regulates telomere length (40). NBS1 colocalizes with TRF1 at PML bodies during late S/G2 phases in immortalized telomerase-negative cells (41) and is encoded by the gene mutated in Nijmegen breakage syndrome, a chromosomal instability disorder. It has been suggested that NBS1 may be involved in alternative lengthening of telomeres in telomerase-negative immortalized cells (41).
TRF1/PIN2 binds telomeres in interphase and will therefore partially colocalize with the chromocenters of mouse nuclei and with SALL1 (22,23). The colocalization between these two proteins in interphase can be expected to be incomplete, since not all telomeres of mouse chromosomes cluster at the chromocenters and the telomeres only occupy a small portion of the chromocenters. At the present time, many questions concerning the exact nature of the interaction of the two proteins and their role for telomere function and/or heterochromatin formation and maintenance remain to be answered. These matters have just become more complicated by the first report of an extra-telomere function and localization of TRF1/PIN2: the protein was localized at the mitotic spindle, suggesting a new role for TRF1/PIN2 in modulating the function of microtubules during mitosis (42). Futhermore, it is tempting to speculate that the interaction between TRF1/PIN2 and SALL1 is involved in the recently reported telomere position effect in human cells, which results in reversible silencing of genes near telomeres depending on the length of the adjacent telomere (43). We have provided strong evidence that besides HP1 (which interacts with Ku70) a second mammalian heterochromatin-associated protein is interacting with a protein involved in telomere function. With these data a picture of a close functional link between the centromeric and telomeric heterochromatin domains not only in Drosophila and yeast, but also in mammalian cells is emerging.
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MATERIALS AND METHODS |
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TOPABSTRACTINTRODUCTIONRESULTSDISCUSSIONMATERIALS AND METHODSREFERENCES |
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Plasmids
Various cDNA fragments of SALL1 were cloned into pBluescript as described previously (44). We subcloned SALL1
1 (bp 1–549 of the complete coding cDNA of SALL1; Fig. 1), SALL12 (bp 261–3175; Fig. 1) and SALL13 (bp 2067–3975; Fig. 1) into the yeast expression vector pGBT9 (Clontech, Palo Alto, CA) by using standard techniques to produce fusion proteins with the GAL4-DNA-binding domain. The construct pGBT9–SALL1 (containing the complete ORF of SALL1; Fig.1) was generated by a multistep subcloning strategy using appropriate restriction enzymes. The full-length SALL1 cDNA was inserted (i) into pEGFP-C1 (CLONTECH) to generate a GFP fusion protein, and (ii) into pM1 (CLONTECH), a mammalian vector used for expression of hybrid proteins with the DNA-binding domain of GAL4. By further subcloning, we generated pM2-SALL12. As a reporter in the transcription repression assays, the plasmid pGAL45tkLUC was used. This plasmid was constructed by replacing the chloramphenicol acetyltransferase gene (CAT) in pGAL45tkCAT (45) with the firefly luciferase gene. This was accomplished by excising the CAT gene from pGAL45tkCAT using BglII and SMI and replacing it by a BglII–SalI fragment containing the luciferase gene from the pGL3-Basic plasmid (Promega, Madison, WI). The resulting construct expresses the luciferase gene under the control of the Herpes simplex virus thymidine kinase promoter.
Cell transfection and fixation
NIH-3T3 cells were grown on 18 x 18 mm coverslips in 5 ml DMEM with 10% FCS and 1% antibiotics at 37°C in a humidified 5% CO2 atmosphere. Transfection with 10 µg pEGFP–SALL1 was carried out with Roti®-Fect (ROTH, Karlsruhe, Germany) according to the manufacturer’s instructions. Forty-eight hours after transfection cells were washed in 1x PBS and fixed by a 10 min incubation in 3.7% formaldehyde (Sigma, Taufkirchen, Germany)/1x PBS at room temperature.
Immunolocalization
For indirect immunofluorescence cells were permeabilized using 0.5% Triton X-100 (Calbiochem, Bad Soden, Germany)/1x PBS for 20 min at room temperature. To avoid unspecific antibody binding, cells were incubated for 15 min at 37°C in a blocking solution containing 4% BSA in PBT [(1x PBS, 0.02% Tween 20 (Calbiochem)]. HP1/M31 detection was accomplished using a commercially available monoclonal rat antibody (Serotec; 10–50 µg/ml) as primary antibody (undiluted) and a Cy3-conjugated goat anti-rat antibody (1 mg/ml; Amersham Pharmacia Biotech, Freiburg, Germany) as secondary antibody (diluted 1:500 in blocking solution). Antibody incubation was performed at 37°C for 30 min. Between the two detection layers, cells were washed in PBT (3 x 3 min at 37°C). Nuclei were counterstained with DAPI (0.05 µg/ml) for epifluorescence microscopy or TO-PRO®-3 (Molecular Probes, Leiden, The Netherlands; 1 µM) for confocal microscopy (DAPI could not be used for confocal imaging as the confocal microscope used was not equipped with a UV-laser. TO-PRO®-3 reveals the same nuclear staining pattern as DAPI). Coverslips were mounted on slides using Vectashield antifade (Vector Laboratories, Burlingame, CA) and sealed with nail polish.
Epifluorescence microscopy
Epifluorescence images were obtained using a Zeiss Axiophot II microscope (Zeiss, Jena, Germany) equipped with a Zeiss Plan-Apochromat 40x/1.4 NA objective lens, and with single-band pass filter sets for visualization of green (GFP), red (Cy3) and infrared (TO-PRO®-3) fluorescence. For image acquisition a cooled charged couple device camera (Photo Science Ltd, Millham, UK) was used; 8 bit grayscale images were recorded and merged to RGB images via Cytovision 2.7 software (Applied Imaging International Ltd, Newcastle-Upon-Tyne, UK). Pictures were processed with Adobe Photoshop 6.0 (Adobe Systems, Mountain View, CA).
Confocal microscopy
Optical sections of 324 or 203 nm were acquired with a Leica TCS SP confocal laser scanning microscope (Leica Microsystems, Heidelberg, Germany) equipped with an oil immersion Plan-Apochromat 100x/1.4 NA objective lens. Fluorochromes were visualized either using an argon laser with excitation wavelengths of 488 nm (for GFP) and 568 nm (for Cy3), or using a helium–neon laser with an excitation wavelength of 633 nm (for TO-PRO®-3). Image resolution was 512 x 512 pixels with a pixel size ranging from 25 to 55 nm depending on the selected zoom factor. RGB pictures were generated by merging confocal images in Adobe Photoshop 6.0.
Transcription repression assay
NIH-3T3 cells (1.4 x 105 cells/35 mm plate) were transfected with Roti®-Fect (ROTH). Each transfection assay included (i) 0.5 µg of the pM1–SALL1 construct, (ii) 0.5 µg of the pGAL45tkLUC reporter plasmid and (iii) 0.1 µg of pCMV-ß-Gal (Clontech), expresssing the ß-galactosidase enzyme. Forty-eight hours after transfection, cells were harvested, lysed in 100 µl of lysis buffer [100 mM KH2PO4 pH 7.8, 0.2% (v/v) Triton X-100, 0.5 mM DTT] and assayed for luciferase and ß-galactosidase activity with an Autolumat LB953 (Berthold, Wildbad, Germany) as described by Schlüter et al. (46). Trichostatin A (TSA) was added to the culture medium after transfection at a final concentration of 500 ng/ml. The levels of ß-galactosidase expression were used to normalize the efficiency of transfection. All assays were repeated at least five times. The median of repression activity and the SD were calculated.
Yeast two-hybrid screen
We used the Matchmaker Two-Hybrid Kit (Clontech) with pGBT9–SALL1-2 as a bait. The assay was performed as recommended by the manufacturer. In brief, yeast cells (strain CG-1945) containing the bait plasmid pGBT9–SALL12 were transformed with a HeLa cDNA library (Clontech) using the lithium acetate method. We screened approximately 2 x 106 transformants for growth on synthetic dropout (SD) plates lacking histidine, leucine, and tryptophan in the presence of 10 mM 3-amino-1,2,4-triazol (3-AT). Colonies growing on the SD plates were analyzed for ß-galactosidase activity by filter assay. The pGAD-GH plasmids containing the cDNA sequence of the putative SALL1 interaction partners were recovered from the yeast cells, transformed into Escherichia coli DH5 and sequenced. The plasmids were then retransformed into the CG-1945 yeast and assayed for growth on SD plates lacking histidine and leucine to exclude DNA-binding activity. In a second round of testing, the pGAD-GH plasmids and pGBT9–SALL12 were cotransformed into CG-1945, and transformants were assayed again for growth on SD plates lacking histidine and for ß-galactosidase activity in a filter assay.
Mapping of interacting domains
To map the SALL1 domains responsible for the interaction with PIN2, yeast strain CG-1945 was cotransformed with SALL1 and its deletion mutants (SALL11, SALL12 and SALL13; Fig. 1) cloned into pGBT9 (Clontech) and PIN2 (Fig. 4) cloned into pGAD-GH (Clontech). Growth was assayed on SD plates lacking leucine and tryptophane (transformation control) or SD plates supplemented with 10 mM 3-AT lacking leucine, tryptophane and histidine (interaction assay). Colonies growing on SD plates lacking leucine, tryptophane and histidine were tested for ß-galactosidase activity in a filter assay.
Glutathion S-transferase (GST)-pulldown and in vitro translation assays
The PIN2-cDNA was cut out of pGAD-GH and cloned into the pGEX-KT vector (Amersham Pharmacia) to produce the fusion protein GST–PIN2 (amino acids 67–331 of PIN2; Fig. 4). Expression and purification of GST–PIN2 and GST alone (as a control) using glutathione Sepharose 4B (Amersham Pharmacia) were performed as recommended by the manufacturer. In vitro translation of SALL12 (amino acids 87–1058 of SALL1; Fig. 1) was performed with rabbit reticulocyte lysate (Promega) and 35S-methionine according to the manufacturer’s instruction. The glutathione Sepharose beads with bound GST fusion proteins (40 µl) and 22 µl of in vitro-translated SALL12 were incubated in 200 µl of bead-binding buffer [50 mM K-phosphate pH 7.5, 100 mM KCl, 10% glycerol (v/v), 0.1% Triton X-100] at 4°C for 2 h. The beads were washed four times with bead-binding buffer devoid of glycerol and Triton X-100. Beads were then heated for 5 min at 95°C in SDS-sample buffer and analyzed by SDS–PAGE. Radiolabeled bands were visualized by autoradiography.
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ACKNOWLEDGEMENTS |
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The authors would like to thank Wolfgang Engel for his support and critically reading the manuscript, Nicole Richter for technical assistance and Folker Garbe for practical help. We are also grateful to Harry Scherthan for helpful discussions and Thomas Cremer for support of the project. This study was funded by the Fritz-Thyssen-Stiftung (grant no. 2000 1071).
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FOOTNOTES |
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+ To whom correspondence should be addressed. Tel: +49 551 397592; Fax: +49 551 399303; Email: cnetzer@web.de. Present addresses: Stefan K.Bohlander and Chang-Dong Zhang, Department of Medicine III, University of Munich, and GSF, Clinical Cooperative Group ‘Leukemia’, Marchioninistrasse 25, 81377 München, Germany
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REFERENCES |
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