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Disruption of the PU.1 gene in chicken B lymphoma DT40 cells and its effect on reported target gene expression
Gene 322 (2003) 169 – 174 www.elsevier.com/locate/gene
Disruption of the PU.1 gene in chicken B lymphoma DT40 cells and its effect on reported target gene expression Hiroki Matsudo, Akira Otsuka, Yukiko Ozawa, Masao Ono * Department of Life Science, and Frontier Project ‘‘Life’s Adaptation Strategies to Environmental Changes’’, College of Science, Rikkyo University, Toshima-ku, Tokyo 171-8501, Japan Received 30 May 2003; received in revised form 11 August 2003; accepted 26 August 2003 Received by T. Sekiya
Abstract Using avian B lymphoma-derived DT40 cells, we disrupted a gene encoding the transcription factor PU.1. The mutant mRNA codes for a protein incapable of functioning as a transcription factor because of the deletion of the protein’s DNA-binding domain. The absence of a functional PU.1 protein in the mutant cells was confirmed by Western blotting and electrophoretic mobility shift assay, thereby demonstrating that PU.1 was not essential for the proliferation of DT40 cells. An examination of the expression of several genes known to be PU.1 protein targets revealed almost the same levels of Ig-h and Ig E L chain mRNA in mutant cells as in wild-type cells, indicating that the PU.1 protein plays no essential role in the transcription of these genes. Mutant cell doubling times 1.3 times longer than those of wild-type cells confirmed the PU.1 protein to be involved in the proliferation of B lymphocytes. D 2003 Elsevier B.V. All rights reserved. Keywords: B lymphocytes; Differentiation; Gene regulation; Transcription factor; Knockout cell
1. Introduction Blood cells are produced via the proliferation and differentiation of pluripotent hematopoietic stem cells possessing self-regenerating activity and the various transcription factors specifically expressed at certain cell lineage and differentiation stages regulate the process. PU.1 is a member of the Ets family of transcription factors and was originally isolated as a protein bound to the purine-rich element 5V-GAGGAA-3Vlocated in the promoter of the murine MHC class II 1-Ab gene (Klemsz et al., 1990). The PU.1 gene is specifically transcribed in B lymphocytes, monocytes, granulocytes, megakaryocytes, mast cells, and immature erythroid cells and is not transcribed in mature T cells (Fisher and Scott, 1998; Simon, 1998; Oikawa et al., 1999). The PU.1 proteins of
Abbreviations: ALV, avian leucosis virus; Bsr, blasticidin; EMSA, electrophoretic mobility shift assay; G3PDH, glyceraldehyde 3-phosphate dehydrogenase; Hyg, hygromycin; mIg, membrane immunoglobulin. * Corresponding author. Tel.: +81-3-3985-2387; fax: +81-3-3985-2387/ 5992-3434. E-mail address: [email protected] (M. Ono). 0378-1119/$ - see front matter D 2003 Elsevier B.V. All rights reserved. doi:10.1016/j.gene.2003.08.021
mouse and human are 272 amino acid residues in length and possess an N-terminal activation domain consisting of a sequence rich in acidic amino acid residues followed by glutamine residues, a central PEST domain that functions in protein degradation and interacts with other transcription factors, and a C-terminal DNA-binding domain (an Ets domain) consisting of a winged helix-turn-helix motif that is rich in basic amino acids (Fisher and Scott, 1998; Simon, 1998; Oikawa et al., 1999. The Ig j, Ig k, Ig l, Ig J chain, Ig-a, Ig-b, CD20, Bruton’s tyrosine kinase, and CD72 genes are known targets of PU.1 in B lymphocytes, whereas the reported targets in myeloid (monocyte/granulocyte) lineages are the G-CSFR, GM-CSFR, M-CSFR, CD11b, CD18, and p47phox (phagocyte NADPH oxidase component) genes (Fisher and Scott, 1998; Simon, 1998; Oikawa et al., 1999). The PU.1 gene is transcribed from multipotential progenitor cells (Chen et al., 1995b) through the plasma cell stage in B cell lineages (Klemsz et al., 1990). Severe immunodeficiency due to defects in B lymphocytes and in myeloid lineages has been reported in mice with a disrupted PU.1 gene (Scott et al., 1994; McKercher et al., 1996). The finding that B cells after the pre-B cell stage
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are absent in PU.1 knockout mice indicates that the PU.1 protein produced at the early stage of B cell differentiation is essential for the differentiation of B cell lineages from the pre-B cell stage onward. The arrest of cytodifferentiation at the stage in which the disrupted gene is essential and the depleted PU.1 protein levels in these knockout mice, however, preclude an elucidation of the gene’s function at this and subsequent stages of differentiation. Consequently, it is not obvious whether PU.1 protein is required for proliferation and differentiation of B lymphocytes at a stage later than the pre-B cell stage. The lack of expression of genes specific to B cells, including known targets of the PU.1 protein such as the Ig-a and Ig-b genes, has been reported in fetal liver cells of PU.1 knockout mice (Scott et al., 1994), but the direct effects of PU.1 protein depletion at later stages of B cell differentiation have not been examined because most PU.1 targets are produced at stages later than the pre-B stage. Part of the explanation for our lack of understanding in this area is that the investigation of target genes of the PU.1 protein have primarily been carried out via electrophoretic mobility shift assay (EMSA), footprint analysis, and transient transfection study. No genetic investigations using PU.1-depleted B lymphocytes have been conducted. DT40 cells are derived from avian leucosis virus (ALV)induced bursal lymphoma. They possess the potential to cause homologous recombination at high frequency and thus are useful in gene disruption studies (Winding and Berchtold, 2001). Ig-h together with membrane immunoglobulin (mIg) and Ig-a constitute the membrane immunoglobulin receptor complex. In the course of our inquiry into the mechanism of B cell-specific transcription of the Ig-b gene in DT40 cells (Katsukura et al., 2001), we discovered an mRNA coding for the PU.1 protein. We have succeeded in disrupting the PU.1 gene and consequently have discovered that the PU.1 protein is not essential for the proliferation of DT40 cells. In this study, the expression of Ig k, Ig l, and Ig-b genes, known targets of PU.1 in B lymphocytes, was examined with PU.1-disrupted DT40 cells.
2. Materials and methods 2.1. Targeting DT40 cells obtained from Dr. Takeda (Kyoto University, Kyoto, Japan) were grown as previously described (Katsukura et al., 2001). A plasmid containing the chicken a-actin promoter-driven hygromycin (Hyg) or blasticidin (Bsr) resistance gene was also obtained from Dr. Takeda. The nucleotide sequence of the cDNA (Klemsz et al., 1990) and organization of the mouse PU.1 gene (McKercher et al., 1996) were used to estimate the exon/intron junctions of chicken PU.1 cDNA (Kherrouche et al., 1998), and the organization of the chicken gene was investigated with polymerase chain reaction (PCR) methods. The chicken PU.1 gene was consequently found to be 7.8 kb in size
Fig. 1. Organization of the chicken PU.1 gene and the structure of the knockout construct (A), examination of target integration by genomic Southern hybridization (B), expression of the PU.1 mRNA in wild-type and double-mutant cells (C), and the amino acid sequence of wild-type and mutant PU.1 proteins (D). (A) Targeting constructs were prepared by the insertions of a 4.2-kb upstream fragment, a resistance gene, and a 1.45-kb downstream fragment into the pCR2.1-TOPO vector. (B) Following digestion of genomic DNA with BamHI and DraI, Southern hybridization was carried out. A 0.3-kb fragment from PU.1 exon 5 (#700 – 998) (Kherrouche et al., 1998) was used as the probe. Lane 1, wild type; lanes 2 and 4, single mutant; lane 3, double mutant. (C) 4 Ag total RNA was separated on a 1% agarose gel containing formaldehyde and transferred to a nitrocellulose filter. The filter was hybridized with the PU.1 cDNA probe. WT, wild-type RNA; DM, double-mutant RNA. The sizes of major bands are shown in the margins. (D) The amino acid sequence was deduced from the nucleotide sequence determined by direct sequencing following RTPCR. Amino acid sequence differences between the wild type and double mutant are indicated by a single-letter code.
(Fig. 1A). The structure of the chicken gene was then used in the design of targeting vectors (Fig. 1A). A 5.2-kb fragment spanning the region from exon 3 to exon 4 was amplified with primers 1F and 2R and cloned into pCR2.1-TOPO (Invitrogen, Carlsbad, CA, USA). Deletion via BamHI digestion
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generated a recombinant plasmid containing a 4.2-kb fragment (a 0.2-kb DNA segment of exon 3 and a 4.0-kb DNA segment of intron 3). A 1.45-kb fragment spanning exons 4 – 5 was amplified with primers 3F and 4R and was cloned into a KpnI site of the recombinant. Finally, the resistance gene was inserted into a BamHI site located in the middle of the upstream and downstream arms (Fig. 1A). The primer sequences correspond to the following nucleotide positions: 1F, #380 –399; 2R, #589– 570; 3F, #580– 599; 4R, #1364 – 1345 (Kherrouche et al., 1998). Both the Hyg and the Bsr genes were inserted in the same orientation as the flanking PU.1 gene. DT40 cells (107) were electroporated with 25 Ag DNA at 550 V and 25 AF in 0.8 ml phosphate-buffered saline with the Gene Pulser II (BioRad Laboratories, Hercules, CA, USA), and the cell suspension was disspensed into 96-well microtiter plates. Clones were obtained in the presence of 1.5 mg/ml Hyg or 30 Ag/ml Bsr. Genomic Southern hybridization with a 0.3-kb probe generated from the PU.1 exon 5 (position #700 –998) (Kherrouche et al., 1998) was used to examine target integration. The preparation of total RNA, formaldehyde/agarose gel electrophoresis, Northern transfer, and hybridization were performed as previously described (Katsukura et al., 2001). The probes used were Ig-b (#313 –1544, 1232 bp; DDBJ/EMBL/GenBank AB062512), Ig l (#873 – 1286, 414 bp; K00389), Ig k (#309 –640, 332 bp; K00678), and Bu-1 (#440 – 1040, 601 bp; X92865). Probe DNA was labeled with [a-32P]-dCTP by the random-priming method. Filter reprobing with human glyceraldehyde 3-phosphate dehydrogenase (G3PDH) cDNA (BD Biosciences Clontech, Palo Alto, CA, USA) was then carried out according to the manufacturer’s instructions. 2.2. Reverse transcriptase-PCR (RT-PCR) and sequencing Using the 3VRACE system (Gibco BRL, Rockville, MD, USA), we synthesized cDNA with an oligo(dT)17-mer primer with a 20-mer adapter and DT40 total RNA. With this cDNA as the template, the first PCR was carried out with primer 5F (#219 – 238 in the exon 1) and AUAP (5VGGCCACGCGTCGACTAGTAC-3V). The second PCR was performed with combinations of the following four primers: 6F, #380– 399 in exon 3; 7F, #504– 523 in exon 3; 8R, #979 – 998 in exon 5; and 9R, #1345 – 1364 in exon 5 (Kherrouche et al., 1998). Amplified DNA was sequenced directly with an ABI PRISM 310 Genetic Analyzer (Applied Biosystems, Foster City, CA, USA). 2.3. Western blotting, electrophoretic mobility shift assay (EMSA), and growth curve determination Western blotting was carried out as described previously (Ono et al., 1994). Antigen detection was performed with the Western Breeze Immunodetection Kit (Invitrogen). A rabbit polyclonal antibody against a peptide mapping at the carboxyl terminus of mouse PU.1 (Santa Cruz Biotechnology, Santa Cruz, CA, USA) was used. The preparation of
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nuclear extracts for EMSA and the end-labeling of the 26mer DNA were performed as previously described (Komatsu et al., 2002). The binding reaction followed by 4.5% polyacrylamide gel electrophoresis was carried out as described (Komatsu et al., 2002). Growth curves were determined by propagating triplicate samples containing 2 104 cells and 2 ml of medium in wells of 6-well plates.
3. Results and discussion 3.1. Targeting Using the targeting vector with the Hyg resistance gene (Fig. 1A), we obtained 13 clones after electroporation and selection. Genomic Southern hybridization with the probe (Fig. 1A), which maps to exon 5 (#700 –988, 298 bp in PU.1 cDNA) (Kherrouche et al., 1998), demonstrated one of the clones to be a disrupted one. To disrupt the other PU.1 gene, we transfected the targeting construct with the Bsr resistance gene into single-mutant cells and obtained 30 Bsrresistant clones. Based on the following procedures, we determined one clone to be a double mutant. DNA digested with BamHI and DraI was prepared from wild-type DT40 cells, and single-mutant cells and putative double-mutant cells were Southern hybridized with the probe. In the DNA prepared from wild-type cells, a 2.3-kb band expected from the gene map was found, whereas in the DNA made from single-mutant cells, a 2.3-kb band from the wild-type allele, and a 1.6-kb band from the mutant allele were detected (Fig. 1B). In DNA prepared from double-mutant cells, only the 1.6-kb band derived from the knockout allele was present (Fig. 1B). 3.2. Expression of PU.1 mRNA in mutant cells and direct sequencing of the mutant PU.1 mRNA Expression of PU.1 mRNA in mutant cells was examined by Northern hybridization. As has been reported previously (Kherrouche et al., 1998), a major 2.1-kb PU.1 mRNA and a minor 2.9-kb mRNA were present in wild-type cells, and a 1.9-kb mRNA was predominantly expressed in the doublemutant cells (Fig. 1C). The nucleotide sequence of the mutant PU.1 mRNA was determined by direct sequencing with the RT-PCR method. Following reverse transcription with total RNA as the template, PU.1 cDNA was amplified by nested PCR. Direct sequencing of the wild-type cDNA demonstrated the presence of an extra 30-bp sequence between exons 3 and 4 (Fig. 1D) (Kherrouche et al., 1998). This region of 10 amino acid residues is coded for by the 5V end of exon 4 in the mouse PU.1 gene (McKercher et al., 1996). The mutant mRNA lacked the extra 30 bp and the 127 bp of exon 4 and thus was 157 bp shorter than the wild-type mRNA. Except for this difference, the nucleotide sequence of the mutant mRNA was otherwise identical to the wild-type mRNA.
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This difference corresponded well with the difference in mRNA sizes (2.1 kb for wild type vs. 1.9 kb for the mutant) detected by Northern hybridization. In the mutant PU.1 mRNA, the connecting of exons 3 and 5 caused a frame shift in exon 5 and generated a TGA stop codon 20 amino acid residues downstream from the 5Vend of exon 5. Thus, the wild-type mRNA was demonstrated to encode a PU.1 protein with 260 amino acid residues, whereas a protein with 123 residues was encoded by the mutant PU.1 mRNA (Fig. 1D). The sequence of the 113 N-terminal residues of the mutant protein was identical to that of the wild-type protein; however, the mutant PU.1 protein lacked the DNAbinding domain encoded in exon 5 (Fisher and Scott, 1998; Oikawa et al., 1999; Simon, 1998) and thus could not function as the transcription factor. The 5Vend of the downstream arm of the targeting construct starts 26 bases downstream from the 5V end of exon 4, and thus the absence of the exon 4 splice acceptor in the disrupted genes is likely to cause a skip of the extra 30 bases and exon 4. 3.3. Absence of the PU.1 protein in mutant cells The absence of the PU.1 protein in mutant cells was demonstrated by Western blotting with an antibody against the C-terminal portion of the mouse PU.1 protein (Fig. 2A). Bands 40 kDa in size that reacted with the antibody were detected in extracts prepared from wild-type cells and single-mutant cells, although the intensity of the band in the single-mutant cells was slightly less than that of wildtype cells. However, no such band was present in doublemutant extracts. Chicken PU.1 has been reported to be 40 kDa in size (Kherrouche et al., 1998), and thus doublemutant cells are shown to lack the protein. Because the antibody used in this study was raised against the C-terminal portion of the protein, our Western blotting procedures would not have detected the 123 N-terminal residues constituting the truncated mutant protein, which may have been present in the mutant cells. 3.4. EMSA analyses of the DNA-binding protein To investigate the DNA binding activity to the PU.1 consensus binding site in the extract prepared from mutant cells, we carried out EMSA analyses (Fig. 2B). As indicated by arrow 1 in this figure, the wild-type extract exhibited a marked gel shift in activity in response to the DNA fragment carrying PU.1 binding site in the promoter of the human cytosolic phagocyte oxidase gene (Li et al., 1999), whereas the mutant extract lacked such an activity, although a faint band that moved slightly faster was detectable. The gel shift activity as indicated by arrow 1 in the wild-type extract competed with a 150-fold excess of the competitor, and the faint band close to arrow 1 observed in the mutant extract was not affected by such competition. In the wild-type extract, no such competition was observed for a competitor with a two-base substitution of GA to CT in the middle of
Fig. 2. Detection of PU.1 protein by Western blotting (A) and the electrophoretic mobility shift assay (EMSA) (B). (A) Total extracts were prepared from wild-type and double-mutant cells, and 20 Ag protein per lane was separated by 12% SDS-polyacrylamide gel electrophoresis. Following the transfer to a nitrocellulose filter, PU.1 protein was detected by the antibody. Lane 1, wild type; lane 2, double mutant; lane 3, single mutant. (B) The binding reaction was carried out with cell extract and labeled DNA1 containing the PU.1 consensus binding site. For the competition experiments, unlabeled DNA1 and its mutant DNA with a twobase substitution in the middle of the consensus binding site were used. Representative shifts 1 and 2 are indicated by arrows. WT, wild type; DM, double mutant.
the consensus binding site. The gel shift bands indicated by arrow 2 were the same in both extracts. Consequently, we concluded that the gel shift indicated by arrow 1 was caused by PU.1 and that the marked decrease in this shift in mutant cells was due to the absence of a PU.1 protein with a DNAbinding domain. The above results indicate that the mutant PU.1 mRNA is incapable of encoding protein with DNA-binding activity, although the level of mutant mRNA was almost the same as that of wild-type mRNA. Even if the mutant PU.1 protein
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with 123 amino acid residues were produced in mutant cells, its lack of a DNA-binding domain precludes its function as a transcription factor. Therefore, we conclude that a PU.1 protein acting as a transcription factor is not essential for the proliferation of bursal lymphoma-derived DT40 cells. 3.5. Expression of the reported target genes in the mutant cells Expression of the reported target genes of PU.1 protein in B cells was examined by Northern hybridization (Fig. 3A). Although a slight increase in expression was observed in mutant cells, no marked differences in the levels of Ig-h and Ig E light chain mRNAs were found in wild-type and mutant cells. The amount of mRNA coding for B cell –specific membrane protein Bu-1 was the same in both cells. In contrast, the levels of Ig A chain mRNA detected in mutant cells were higher than in wild-type cells. The presence of PU.1 binding sites is indicated in the promoter of Ig-b gene
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(Omori and Wall, 1993; Thompson et al., 1996), in the 3Venhancer of the Ig k gene (Eisenbeis et al., 1993; Eisenbeis et al., 1995), and in the intron enhancer of the Ig l gene (Nelsen et al., 1993; Rivera et al., 1993). In addition, reporter assays demonstrated these sites to be involved in the expression of the target gene. However, there were no large differences in the levels of Ig-h and Ig E mRNAs expressed in wild-type and mutant cells, indicating that PU.1 does not play an essential role in the transcription of these genes in DT40 cells. The absence of Ig-h mRNA in fetal liver cells of PU.1 knockout mice does not necessarily indicate that the PU.1 is essential for the transcription of the Ig-b gene, because the knockout itself may be responsible for the absence of cells producing Ig-h mRNA. The PU.1 protein has been reported to participate in the transcription of its own gene (Chen et al., 1995a); however, the amounts of wild-type and mutant mRNA in the present study were almost the same (Fig. 1C), so the PU.1 protein cannot have an essential role in PU.1 gene transcription. The reason for the inconsistency in the results of the present study and previous ones may derive from the different methods used. Earlier studies were based on factor binding and transient transfection assays, whereas this study used a genetic approach. Difference in mammals and chickens may be another reason because no such assay has been carried out in chickens. Since we only compared the steady-state mRNA levels of the reported target genes by Northern hybridization, it is difficult to conclude that PU.1 protein is not involved in the regulation of Ig-h and Ig E genes. To make these points clear, promoter – reporter assays should be carried out. The observed up-regulation of Ig A mRNA expression in mutant cells was opposite to the other reported results. Although, again, the reason for this difference remains to be resolved, it is now less likely that PU.1 acts as a positive regulator for Ig l gene transcription. 3.6. Growth characteristics of mutant cells
Fig. 3. Expression of the reported target genes for PU.1 (A) and the growth characteristics of double-mutant cells (B). (A) Total RNA (3 Ag) was separated on a 1% agarose gel containing formaldehyde and Northern hybridization was carried out as described in the legend to Fig. 1C. WT, wild type; DM, double mutant. Reprobing with human glyceraldehyde 3phosphate dehydrogenase (G3PDH) cDNA is shown at the bottom. (B) Wild-type, single-mutant and double-mutant cells were propagated in sixwell plates. Open circle, wild type; open square, single mutant; closed circle, double mutant. Bars indicate standard error.
An examination of growth rates revealed doubling times of wild-type cells and single-mutant cells to be the same (9 h), whereas the doubling time of double-mutant cells was 12 h, 1.3 times longer than that of wild type (Fig. 3B). The densities of all three kinds of cells were the same (6 106/ ml). The fact that PU.1 is identical to a protein encoded by the oncogene Spi-1, which is activated by the insertion of a provirus in Friend virus-induced murine erythroleukemia (Moreau-Gachelin, 1994), indicates that PU.1 is involved not only in the differentiation but also in the proliferation of hematopoietic cells. Studies using cultured mammalian cells have demonstrated that the effect of PU.1 on growth and differentiation in the course of erythroid and myeloid differentiation is different, depending on the cell lineage and the level of expression, and thus is complex (Fisher and Scott, 1998). In contrast, no investigation of the relationship of PU.1 to cell proliferation in B cells has been carried out. This study has demonstrated the involvement of PU.1 in the
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proliferation of DT40 cells, because the mutant cells exhibited a doubling time that was 1.3 times longer. In evaluating the role of PU.1 function in the process of cell proliferation, it is of interest to discover when in the cell cycle the increase in doubling time occurs.
Acknowledgements We thank Dr. S. Takeda for providing DT40 cells and plasmids for the targeting construct. This work was supported by Rikkyo University for the Promotion of Research.
References Chen, H., Ray-Gallet, D., Zhang, P., Hetherington, C.J., Gonzalez, D.A., Zhang, D.E., Moreau-Gachelin, F., Tenen, D.G., 1995a. PU.1 (Spi-1) autoregulates its expression in myeloid cells. Oncogene 11, 1549 – 1560. Chen, H.M., Zhang, P., Voso, M.T., Hohaus, S., Gonzalez, D.A., Glass, C.K., Zhang, D.E., Tenen, D.G., 1995b. Neutrophils and monocytes express high levels of PU.1 (Spi-1) but not Spi-B. Blood 85, 2918 – 2928. Eisenbeis, C.F., Singh, H., Storb, U., 1993. PU.1 is a component of a multiprotein complex which binds an essential site in the murine immunoglobulin lambda 2 – 4 enhancer. Mol. Cell. Biol. 13, 6452 – 6461. Eisenbeis, C.F., Singh, H., Storb, U., 1995. Pip, a novel IRF family member, is a lymphoid-specific, PU.1-dependent transcriptional activator. Genes Dev. 9, 1377 – 1387. Fisher, R.C., Scott, E.W., 1998. Role of PU.1 in hematopoiesis. Stem Cells 16, 25 – 37. Katsukura, H., Murakami, R., Chijiiwa, Y., Otsuka, A., Tanaka, M., Nakashima, K., Ono, M., 2001. Structure of the beta-chain (B29) gene of the chicken B-cell receptor and conserved collinearity with genes for potential skeletal muscle sodium channel and growth hormone. Immunogenetics 53, 770 – 775. Kherrouche, Z., Beuscart, A., Huguet, C., Flourens, A., Moreau-Gachelin, F., Stehelin, D., Coll, J., 1998. Isolation and characterization of a chicken homologue of the Spi-1/PU.1 transcription factor. Oncogene 16, 1357 – 1367.
Klemsz, M.J., McKercher, S.R., Celada, A., Van Beveren, C., Maki, R.A., 1990. The macrophage and B cell-specific transcription factor PU.1 is related to the ets oncogene. Cell 61, 113 – 124. Komatsu, A., Otsuka, A., Ono, M., 2002. Novel regulatory regions found downstream of the rat B29/Ig-beta gene. Eur. J. Biochem. 269, 1227 – 1236. Li, S.L., Schlegel, W., Valente, A.J., Clark, R.A., 1999. Critical flanking sequences of PU.1 binding sites in myeloid-specific promoters. J. Biol. Chem. 274, 32453 – 32460. McKercher, S.R., Torbett, B.E., Anderson, K.L., Henkel, G.W., Vestal, D.J., Baribault, H., Klemsz, M., Feeney, A.J., Wu, G.E., Paige, C.J., Maki, R.A., 1996. Targeted disruption of the PU.1 gene results in multiple hematopoietic abnormalities. EMBO J. 15, 5647 – 5658. Moreau-Gachelin, F., 1994. Spi-1/PU.1: an oncogene of the Ets family. Biochim. Biophys. Acta 1198, 149 – 163. Nelsen, B., Tian, G., Erman, B., Gregoire, J., Maki, R., Graves, B., Sen, R., 1993. Regulation of lymphoid-specific immunoglobulin mu heavy chain gene enhancer by ETS-domain proteins. Science 261, 82 – 86. Oikawa, T., Yamada, T., Kihara-Negishi, F., Yamamoto, H., Kondoh, N., Hitomi, Y., Hashimoto, Y., 1999. The role of Ets family transcription factor PU.1 in hematopoietic cell differentiation, proliferation and apoptosis. Cell Death Differ. 6, 599 – 608. Omori, S.A., Wall, R., 1993. Multiple motifs regulate the B-cell-specific promoter of the B29 gene. Proc. Natl. Acad. Sci. U. S. A. 90, 11723 – 11727. Ono, M., Harigai, T., Kaneko, T., Sato, Y., Ihara, S., Kawauchi, H., 1994. Pit-1/GH factor-1 involvement in the gene expression of somatolactin. Mol. Endocrinol. 8, 109 – 115. Rivera, R.R., Stuiver, M.H., Steenbergen, R., Murre, C., 1993. Ets proteins: new factors that regulate immunoglobulin heavy-chain gene expression. Mol. Cell. Biol. 13, 7163 – 7169. Scott, E.W., Simon, M.C., Anastasi, J., Singh, H., 1994. Requirement of transcription factor PU.1 in the development of multiple hematopoietic lineages. Science 265, 1573 – 1577. Simon, M.C., 1998. PU.1 and hematopoiesis: lessons learned from gene targeting experiments. Semin. Immunol. 10, 111 – 118. Thompson, A.A., Wood Jr., W.J., Gilly, M.J., Damore, M.A., Omori, S.A., Wall, R., 1996. The promoter and 5Vflanking sequences controlling human B29 gene expression. Blood 87, 666 – 673. Winding, P., Berchtold, M.W., 2001. The chicken B cell line DT40: a novel tool for gene disruption experiments. J. Immunol. Methods 249, 1 – 16.
Disruption of the PU.1 gene in chicken B lymphoma DT40 cells and its effect on reported target gene expression Hiroki Matsudo, Akira Otsuka, Yukiko Ozawa, Masao Ono * Department of Life Science, and Frontier Project ‘‘Life’s Adaptation Strategies to Environmental Changes’’, College of Science, Rikkyo University, Toshima-ku, Tokyo 171-8501, Japan Received 30 May 2003; received in revised form 11 August 2003; accepted 26 August 2003 Received by T. Sekiya
Abstract Using avian B lymphoma-derived DT40 cells, we disrupted a gene encoding the transcription factor PU.1. The mutant mRNA codes for a protein incapable of functioning as a transcription factor because of the deletion of the protein’s DNA-binding domain. The absence of a functional PU.1 protein in the mutant cells was confirmed by Western blotting and electrophoretic mobility shift assay, thereby demonstrating that PU.1 was not essential for the proliferation of DT40 cells. An examination of the expression of several genes known to be PU.1 protein targets revealed almost the same levels of Ig-h and Ig E L chain mRNA in mutant cells as in wild-type cells, indicating that the PU.1 protein plays no essential role in the transcription of these genes. Mutant cell doubling times 1.3 times longer than those of wild-type cells confirmed the PU.1 protein to be involved in the proliferation of B lymphocytes. D 2003 Elsevier B.V. All rights reserved. Keywords: B lymphocytes; Differentiation; Gene regulation; Transcription factor; Knockout cell
1. Introduction Blood cells are produced via the proliferation and differentiation of pluripotent hematopoietic stem cells possessing self-regenerating activity and the various transcription factors specifically expressed at certain cell lineage and differentiation stages regulate the process. PU.1 is a member of the Ets family of transcription factors and was originally isolated as a protein bound to the purine-rich element 5V-GAGGAA-3Vlocated in the promoter of the murine MHC class II 1-Ab gene (Klemsz et al., 1990). The PU.1 gene is specifically transcribed in B lymphocytes, monocytes, granulocytes, megakaryocytes, mast cells, and immature erythroid cells and is not transcribed in mature T cells (Fisher and Scott, 1998; Simon, 1998; Oikawa et al., 1999). The PU.1 proteins of
Abbreviations: ALV, avian leucosis virus; Bsr, blasticidin; EMSA, electrophoretic mobility shift assay; G3PDH, glyceraldehyde 3-phosphate dehydrogenase; Hyg, hygromycin; mIg, membrane immunoglobulin. * Corresponding author. Tel.: +81-3-3985-2387; fax: +81-3-3985-2387/ 5992-3434. E-mail address: [email protected] (M. Ono). 0378-1119/$ - see front matter D 2003 Elsevier B.V. All rights reserved. doi:10.1016/j.gene.2003.08.021
mouse and human are 272 amino acid residues in length and possess an N-terminal activation domain consisting of a sequence rich in acidic amino acid residues followed by glutamine residues, a central PEST domain that functions in protein degradation and interacts with other transcription factors, and a C-terminal DNA-binding domain (an Ets domain) consisting of a winged helix-turn-helix motif that is rich in basic amino acids (Fisher and Scott, 1998; Simon, 1998; Oikawa et al., 1999. The Ig j, Ig k, Ig l, Ig J chain, Ig-a, Ig-b, CD20, Bruton’s tyrosine kinase, and CD72 genes are known targets of PU.1 in B lymphocytes, whereas the reported targets in myeloid (monocyte/granulocyte) lineages are the G-CSFR, GM-CSFR, M-CSFR, CD11b, CD18, and p47phox (phagocyte NADPH oxidase component) genes (Fisher and Scott, 1998; Simon, 1998; Oikawa et al., 1999). The PU.1 gene is transcribed from multipotential progenitor cells (Chen et al., 1995b) through the plasma cell stage in B cell lineages (Klemsz et al., 1990). Severe immunodeficiency due to defects in B lymphocytes and in myeloid lineages has been reported in mice with a disrupted PU.1 gene (Scott et al., 1994; McKercher et al., 1996). The finding that B cells after the pre-B cell stage
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are absent in PU.1 knockout mice indicates that the PU.1 protein produced at the early stage of B cell differentiation is essential for the differentiation of B cell lineages from the pre-B cell stage onward. The arrest of cytodifferentiation at the stage in which the disrupted gene is essential and the depleted PU.1 protein levels in these knockout mice, however, preclude an elucidation of the gene’s function at this and subsequent stages of differentiation. Consequently, it is not obvious whether PU.1 protein is required for proliferation and differentiation of B lymphocytes at a stage later than the pre-B cell stage. The lack of expression of genes specific to B cells, including known targets of the PU.1 protein such as the Ig-a and Ig-b genes, has been reported in fetal liver cells of PU.1 knockout mice (Scott et al., 1994), but the direct effects of PU.1 protein depletion at later stages of B cell differentiation have not been examined because most PU.1 targets are produced at stages later than the pre-B stage. Part of the explanation for our lack of understanding in this area is that the investigation of target genes of the PU.1 protein have primarily been carried out via electrophoretic mobility shift assay (EMSA), footprint analysis, and transient transfection study. No genetic investigations using PU.1-depleted B lymphocytes have been conducted. DT40 cells are derived from avian leucosis virus (ALV)induced bursal lymphoma. They possess the potential to cause homologous recombination at high frequency and thus are useful in gene disruption studies (Winding and Berchtold, 2001). Ig-h together with membrane immunoglobulin (mIg) and Ig-a constitute the membrane immunoglobulin receptor complex. In the course of our inquiry into the mechanism of B cell-specific transcription of the Ig-b gene in DT40 cells (Katsukura et al., 2001), we discovered an mRNA coding for the PU.1 protein. We have succeeded in disrupting the PU.1 gene and consequently have discovered that the PU.1 protein is not essential for the proliferation of DT40 cells. In this study, the expression of Ig k, Ig l, and Ig-b genes, known targets of PU.1 in B lymphocytes, was examined with PU.1-disrupted DT40 cells.
2. Materials and methods 2.1. Targeting DT40 cells obtained from Dr. Takeda (Kyoto University, Kyoto, Japan) were grown as previously described (Katsukura et al., 2001). A plasmid containing the chicken a-actin promoter-driven hygromycin (Hyg) or blasticidin (Bsr) resistance gene was also obtained from Dr. Takeda. The nucleotide sequence of the cDNA (Klemsz et al., 1990) and organization of the mouse PU.1 gene (McKercher et al., 1996) were used to estimate the exon/intron junctions of chicken PU.1 cDNA (Kherrouche et al., 1998), and the organization of the chicken gene was investigated with polymerase chain reaction (PCR) methods. The chicken PU.1 gene was consequently found to be 7.8 kb in size
Fig. 1. Organization of the chicken PU.1 gene and the structure of the knockout construct (A), examination of target integration by genomic Southern hybridization (B), expression of the PU.1 mRNA in wild-type and double-mutant cells (C), and the amino acid sequence of wild-type and mutant PU.1 proteins (D). (A) Targeting constructs were prepared by the insertions of a 4.2-kb upstream fragment, a resistance gene, and a 1.45-kb downstream fragment into the pCR2.1-TOPO vector. (B) Following digestion of genomic DNA with BamHI and DraI, Southern hybridization was carried out. A 0.3-kb fragment from PU.1 exon 5 (#700 – 998) (Kherrouche et al., 1998) was used as the probe. Lane 1, wild type; lanes 2 and 4, single mutant; lane 3, double mutant. (C) 4 Ag total RNA was separated on a 1% agarose gel containing formaldehyde and transferred to a nitrocellulose filter. The filter was hybridized with the PU.1 cDNA probe. WT, wild-type RNA; DM, double-mutant RNA. The sizes of major bands are shown in the margins. (D) The amino acid sequence was deduced from the nucleotide sequence determined by direct sequencing following RTPCR. Amino acid sequence differences between the wild type and double mutant are indicated by a single-letter code.
(Fig. 1A). The structure of the chicken gene was then used in the design of targeting vectors (Fig. 1A). A 5.2-kb fragment spanning the region from exon 3 to exon 4 was amplified with primers 1F and 2R and cloned into pCR2.1-TOPO (Invitrogen, Carlsbad, CA, USA). Deletion via BamHI digestion
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generated a recombinant plasmid containing a 4.2-kb fragment (a 0.2-kb DNA segment of exon 3 and a 4.0-kb DNA segment of intron 3). A 1.45-kb fragment spanning exons 4 – 5 was amplified with primers 3F and 4R and was cloned into a KpnI site of the recombinant. Finally, the resistance gene was inserted into a BamHI site located in the middle of the upstream and downstream arms (Fig. 1A). The primer sequences correspond to the following nucleotide positions: 1F, #380 –399; 2R, #589– 570; 3F, #580– 599; 4R, #1364 – 1345 (Kherrouche et al., 1998). Both the Hyg and the Bsr genes were inserted in the same orientation as the flanking PU.1 gene. DT40 cells (107) were electroporated with 25 Ag DNA at 550 V and 25 AF in 0.8 ml phosphate-buffered saline with the Gene Pulser II (BioRad Laboratories, Hercules, CA, USA), and the cell suspension was disspensed into 96-well microtiter plates. Clones were obtained in the presence of 1.5 mg/ml Hyg or 30 Ag/ml Bsr. Genomic Southern hybridization with a 0.3-kb probe generated from the PU.1 exon 5 (position #700 –998) (Kherrouche et al., 1998) was used to examine target integration. The preparation of total RNA, formaldehyde/agarose gel electrophoresis, Northern transfer, and hybridization were performed as previously described (Katsukura et al., 2001). The probes used were Ig-b (#313 –1544, 1232 bp; DDBJ/EMBL/GenBank AB062512), Ig l (#873 – 1286, 414 bp; K00389), Ig k (#309 –640, 332 bp; K00678), and Bu-1 (#440 – 1040, 601 bp; X92865). Probe DNA was labeled with [a-32P]-dCTP by the random-priming method. Filter reprobing with human glyceraldehyde 3-phosphate dehydrogenase (G3PDH) cDNA (BD Biosciences Clontech, Palo Alto, CA, USA) was then carried out according to the manufacturer’s instructions. 2.2. Reverse transcriptase-PCR (RT-PCR) and sequencing Using the 3VRACE system (Gibco BRL, Rockville, MD, USA), we synthesized cDNA with an oligo(dT)17-mer primer with a 20-mer adapter and DT40 total RNA. With this cDNA as the template, the first PCR was carried out with primer 5F (#219 – 238 in the exon 1) and AUAP (5VGGCCACGCGTCGACTAGTAC-3V). The second PCR was performed with combinations of the following four primers: 6F, #380– 399 in exon 3; 7F, #504– 523 in exon 3; 8R, #979 – 998 in exon 5; and 9R, #1345 – 1364 in exon 5 (Kherrouche et al., 1998). Amplified DNA was sequenced directly with an ABI PRISM 310 Genetic Analyzer (Applied Biosystems, Foster City, CA, USA). 2.3. Western blotting, electrophoretic mobility shift assay (EMSA), and growth curve determination Western blotting was carried out as described previously (Ono et al., 1994). Antigen detection was performed with the Western Breeze Immunodetection Kit (Invitrogen). A rabbit polyclonal antibody against a peptide mapping at the carboxyl terminus of mouse PU.1 (Santa Cruz Biotechnology, Santa Cruz, CA, USA) was used. The preparation of
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nuclear extracts for EMSA and the end-labeling of the 26mer DNA were performed as previously described (Komatsu et al., 2002). The binding reaction followed by 4.5% polyacrylamide gel electrophoresis was carried out as described (Komatsu et al., 2002). Growth curves were determined by propagating triplicate samples containing 2 104 cells and 2 ml of medium in wells of 6-well plates.
3. Results and discussion 3.1. Targeting Using the targeting vector with the Hyg resistance gene (Fig. 1A), we obtained 13 clones after electroporation and selection. Genomic Southern hybridization with the probe (Fig. 1A), which maps to exon 5 (#700 –988, 298 bp in PU.1 cDNA) (Kherrouche et al., 1998), demonstrated one of the clones to be a disrupted one. To disrupt the other PU.1 gene, we transfected the targeting construct with the Bsr resistance gene into single-mutant cells and obtained 30 Bsrresistant clones. Based on the following procedures, we determined one clone to be a double mutant. DNA digested with BamHI and DraI was prepared from wild-type DT40 cells, and single-mutant cells and putative double-mutant cells were Southern hybridized with the probe. In the DNA prepared from wild-type cells, a 2.3-kb band expected from the gene map was found, whereas in the DNA made from single-mutant cells, a 2.3-kb band from the wild-type allele, and a 1.6-kb band from the mutant allele were detected (Fig. 1B). In DNA prepared from double-mutant cells, only the 1.6-kb band derived from the knockout allele was present (Fig. 1B). 3.2. Expression of PU.1 mRNA in mutant cells and direct sequencing of the mutant PU.1 mRNA Expression of PU.1 mRNA in mutant cells was examined by Northern hybridization. As has been reported previously (Kherrouche et al., 1998), a major 2.1-kb PU.1 mRNA and a minor 2.9-kb mRNA were present in wild-type cells, and a 1.9-kb mRNA was predominantly expressed in the doublemutant cells (Fig. 1C). The nucleotide sequence of the mutant PU.1 mRNA was determined by direct sequencing with the RT-PCR method. Following reverse transcription with total RNA as the template, PU.1 cDNA was amplified by nested PCR. Direct sequencing of the wild-type cDNA demonstrated the presence of an extra 30-bp sequence between exons 3 and 4 (Fig. 1D) (Kherrouche et al., 1998). This region of 10 amino acid residues is coded for by the 5V end of exon 4 in the mouse PU.1 gene (McKercher et al., 1996). The mutant mRNA lacked the extra 30 bp and the 127 bp of exon 4 and thus was 157 bp shorter than the wild-type mRNA. Except for this difference, the nucleotide sequence of the mutant mRNA was otherwise identical to the wild-type mRNA.
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This difference corresponded well with the difference in mRNA sizes (2.1 kb for wild type vs. 1.9 kb for the mutant) detected by Northern hybridization. In the mutant PU.1 mRNA, the connecting of exons 3 and 5 caused a frame shift in exon 5 and generated a TGA stop codon 20 amino acid residues downstream from the 5Vend of exon 5. Thus, the wild-type mRNA was demonstrated to encode a PU.1 protein with 260 amino acid residues, whereas a protein with 123 residues was encoded by the mutant PU.1 mRNA (Fig. 1D). The sequence of the 113 N-terminal residues of the mutant protein was identical to that of the wild-type protein; however, the mutant PU.1 protein lacked the DNAbinding domain encoded in exon 5 (Fisher and Scott, 1998; Oikawa et al., 1999; Simon, 1998) and thus could not function as the transcription factor. The 5Vend of the downstream arm of the targeting construct starts 26 bases downstream from the 5V end of exon 4, and thus the absence of the exon 4 splice acceptor in the disrupted genes is likely to cause a skip of the extra 30 bases and exon 4. 3.3. Absence of the PU.1 protein in mutant cells The absence of the PU.1 protein in mutant cells was demonstrated by Western blotting with an antibody against the C-terminal portion of the mouse PU.1 protein (Fig. 2A). Bands 40 kDa in size that reacted with the antibody were detected in extracts prepared from wild-type cells and single-mutant cells, although the intensity of the band in the single-mutant cells was slightly less than that of wildtype cells. However, no such band was present in doublemutant extracts. Chicken PU.1 has been reported to be 40 kDa in size (Kherrouche et al., 1998), and thus doublemutant cells are shown to lack the protein. Because the antibody used in this study was raised against the C-terminal portion of the protein, our Western blotting procedures would not have detected the 123 N-terminal residues constituting the truncated mutant protein, which may have been present in the mutant cells. 3.4. EMSA analyses of the DNA-binding protein To investigate the DNA binding activity to the PU.1 consensus binding site in the extract prepared from mutant cells, we carried out EMSA analyses (Fig. 2B). As indicated by arrow 1 in this figure, the wild-type extract exhibited a marked gel shift in activity in response to the DNA fragment carrying PU.1 binding site in the promoter of the human cytosolic phagocyte oxidase gene (Li et al., 1999), whereas the mutant extract lacked such an activity, although a faint band that moved slightly faster was detectable. The gel shift activity as indicated by arrow 1 in the wild-type extract competed with a 150-fold excess of the competitor, and the faint band close to arrow 1 observed in the mutant extract was not affected by such competition. In the wild-type extract, no such competition was observed for a competitor with a two-base substitution of GA to CT in the middle of
Fig. 2. Detection of PU.1 protein by Western blotting (A) and the electrophoretic mobility shift assay (EMSA) (B). (A) Total extracts were prepared from wild-type and double-mutant cells, and 20 Ag protein per lane was separated by 12% SDS-polyacrylamide gel electrophoresis. Following the transfer to a nitrocellulose filter, PU.1 protein was detected by the antibody. Lane 1, wild type; lane 2, double mutant; lane 3, single mutant. (B) The binding reaction was carried out with cell extract and labeled DNA1 containing the PU.1 consensus binding site. For the competition experiments, unlabeled DNA1 and its mutant DNA with a twobase substitution in the middle of the consensus binding site were used. Representative shifts 1 and 2 are indicated by arrows. WT, wild type; DM, double mutant.
the consensus binding site. The gel shift bands indicated by arrow 2 were the same in both extracts. Consequently, we concluded that the gel shift indicated by arrow 1 was caused by PU.1 and that the marked decrease in this shift in mutant cells was due to the absence of a PU.1 protein with a DNAbinding domain. The above results indicate that the mutant PU.1 mRNA is incapable of encoding protein with DNA-binding activity, although the level of mutant mRNA was almost the same as that of wild-type mRNA. Even if the mutant PU.1 protein
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with 123 amino acid residues were produced in mutant cells, its lack of a DNA-binding domain precludes its function as a transcription factor. Therefore, we conclude that a PU.1 protein acting as a transcription factor is not essential for the proliferation of bursal lymphoma-derived DT40 cells. 3.5. Expression of the reported target genes in the mutant cells Expression of the reported target genes of PU.1 protein in B cells was examined by Northern hybridization (Fig. 3A). Although a slight increase in expression was observed in mutant cells, no marked differences in the levels of Ig-h and Ig E light chain mRNAs were found in wild-type and mutant cells. The amount of mRNA coding for B cell –specific membrane protein Bu-1 was the same in both cells. In contrast, the levels of Ig A chain mRNA detected in mutant cells were higher than in wild-type cells. The presence of PU.1 binding sites is indicated in the promoter of Ig-b gene
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(Omori and Wall, 1993; Thompson et al., 1996), in the 3Venhancer of the Ig k gene (Eisenbeis et al., 1993; Eisenbeis et al., 1995), and in the intron enhancer of the Ig l gene (Nelsen et al., 1993; Rivera et al., 1993). In addition, reporter assays demonstrated these sites to be involved in the expression of the target gene. However, there were no large differences in the levels of Ig-h and Ig E mRNAs expressed in wild-type and mutant cells, indicating that PU.1 does not play an essential role in the transcription of these genes in DT40 cells. The absence of Ig-h mRNA in fetal liver cells of PU.1 knockout mice does not necessarily indicate that the PU.1 is essential for the transcription of the Ig-b gene, because the knockout itself may be responsible for the absence of cells producing Ig-h mRNA. The PU.1 protein has been reported to participate in the transcription of its own gene (Chen et al., 1995a); however, the amounts of wild-type and mutant mRNA in the present study were almost the same (Fig. 1C), so the PU.1 protein cannot have an essential role in PU.1 gene transcription. The reason for the inconsistency in the results of the present study and previous ones may derive from the different methods used. Earlier studies were based on factor binding and transient transfection assays, whereas this study used a genetic approach. Difference in mammals and chickens may be another reason because no such assay has been carried out in chickens. Since we only compared the steady-state mRNA levels of the reported target genes by Northern hybridization, it is difficult to conclude that PU.1 protein is not involved in the regulation of Ig-h and Ig E genes. To make these points clear, promoter – reporter assays should be carried out. The observed up-regulation of Ig A mRNA expression in mutant cells was opposite to the other reported results. Although, again, the reason for this difference remains to be resolved, it is now less likely that PU.1 acts as a positive regulator for Ig l gene transcription. 3.6. Growth characteristics of mutant cells
Fig. 3. Expression of the reported target genes for PU.1 (A) and the growth characteristics of double-mutant cells (B). (A) Total RNA (3 Ag) was separated on a 1% agarose gel containing formaldehyde and Northern hybridization was carried out as described in the legend to Fig. 1C. WT, wild type; DM, double mutant. Reprobing with human glyceraldehyde 3phosphate dehydrogenase (G3PDH) cDNA is shown at the bottom. (B) Wild-type, single-mutant and double-mutant cells were propagated in sixwell plates. Open circle, wild type; open square, single mutant; closed circle, double mutant. Bars indicate standard error.
An examination of growth rates revealed doubling times of wild-type cells and single-mutant cells to be the same (9 h), whereas the doubling time of double-mutant cells was 12 h, 1.3 times longer than that of wild type (Fig. 3B). The densities of all three kinds of cells were the same (6 106/ ml). The fact that PU.1 is identical to a protein encoded by the oncogene Spi-1, which is activated by the insertion of a provirus in Friend virus-induced murine erythroleukemia (Moreau-Gachelin, 1994), indicates that PU.1 is involved not only in the differentiation but also in the proliferation of hematopoietic cells. Studies using cultured mammalian cells have demonstrated that the effect of PU.1 on growth and differentiation in the course of erythroid and myeloid differentiation is different, depending on the cell lineage and the level of expression, and thus is complex (Fisher and Scott, 1998). In contrast, no investigation of the relationship of PU.1 to cell proliferation in B cells has been carried out. This study has demonstrated the involvement of PU.1 in the
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proliferation of DT40 cells, because the mutant cells exhibited a doubling time that was 1.3 times longer. In evaluating the role of PU.1 function in the process of cell proliferation, it is of interest to discover when in the cell cycle the increase in doubling time occurs.
Acknowledgements We thank Dr. S. Takeda for providing DT40 cells and plasmids for the targeting construct. This work was supported by Rikkyo University for the Promotion of Research.
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