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Genes to Cells (2005) 10, 139-150. doi:10.1111/j.1365-2443.2005.00825.x
© 2005 Blackwell Publishing or its licensors
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Identification of DRG family regulatory proteins (DFRPs): specific regulation of DRG1 and DRG2

Kosuke Ishikawa1, Sakura Azuma1, Shuntaro Ikawa2, Kentaro Semba1 and Jun-ichiro Inoue1,*

1 Division of Cellular and Molecular Biology, Department of Cancer Biology, Institute of Medical Science, The University of Tokyo, 4-6-1 Shirokanedai, Minatoku, Tokyo 108-8639, Japan
2 Department of Cell Biology, Institute of Development, Ageing and Cancer, Tohoku University, Seiryo-cho, Aobaku, Sendai City, Miyagi 980-8575, Japan


   Abstract
Top
 Abstract
Introduction
 Results
 Discussion
 Experimental procedures
 References
 
DRG1 and DRG2 comprise a highly conserved subfamily of GTP-binding proteins and are thought to act as critical regulators of cell growth. Their abnormal expressions may trigger cell transformation or cell cycle arrest. Our aim is to clarify their physiological functions and regulatory mechanisms. Here we report identification of novel proteins, DRG family regulatory protein (DFRP) 1 and DFRP2, which regulate expression of DRG proteins through specific binding. In transient transfection experiments, DFRP1 specifically binds DRG1, and DFRP2 preferentially binds DRG2. DFRPs provide stability to the target DRG proteins through physical association, possibly by blocking the poly-ubiquitination that would precede proteolysis of DRG proteins. DFRPs are highly conserved in eucaryotes, and the expression patterns of dfrp1 and drg1 transcripts in Xenopus embryos and tissues are similar, indicating that these genes work cooperatively in various types of eukaryotic cells. Immunofluorescence experiments have revealed that the interaction between DRG1 and DFRP1 may occur in the cytoplasm. We generated dfrp1- knockout cells and found that endogenous expression of DRG1 is regulated by DFRP1, confirming that DFRP1 is a specific up-regulator of DRG1 in vivo. On the basis of these results, we propose that DRG1 and DRG2 are regulated differently despite their structural similarities.


   Introduction
Top
 Abstract
 Introduction
Results
 Discussion
 Experimental procedures
 References
 
The developmentally regulated GTP-binding protein (DRG) subfamily constitutes one branch of the GTPase superfamily (Leipe et al. 2002). drg was originally identified by a subtractive cDNA cloning approach as a gene (NEDD-3) that is expressed at high levels in developing mouse brain (Kumar et al. 1992; Sazuka et al. 1992a,b). Genes homologous to mouse drg have been reported in a wide variety of eukaryotic and archaean species. These DRG proteins show striking similarities to each other, suggesting that the cellular roles of DRGs may be very important. Schenker & Trueb (1997) carried out a DRG sequence search and reported that the DRG subfamily contains two closely related proteins, DRG1 (for original DRG) and DRG2. Detailed computational analysis with the use of existing genomic databases revealed that eucaryotes have both drg1 and drg2 genes, whereas archaea carry only one drg gene, and it does not specifically correspond to drg1 or drg2 (Li & Trueb 2000; our unpublished observation). Arabidopis thaliana drg2 is highly expressed in actively growing tissues and reproductive organs (Devitt et al. 1999; Etheridge et al. 1999). Human DRG1 associates with SCL (TAL-1) oncogenic protein and stimulates the co-transforming activity of c-Myc and Ras (Mahajan et al. 1996). Over-expression of human DRG2 causes cell cycle arrest at G2/M phase (Ko et al. 2004; Song et al. 2004). These results suggest that DRG proteins play a critical role in cell growth and that their aberrant expression triggers the disruption of normal growth control. Therefore, clarification of physiological functions of DRG proteins and identification of their regulatory molecules may lead to the discovery of a novel cellular mechanism underlying cell growth.

DRG1 and DRG2 are highly homologous across their entire lengths. Regions showing particularly strong homology include a domain of characteristic G-motifs that may be the core of GTPase activity as well as the C-terminal TGS domain, which is related to RNA binding (Ishikawa et al. 2003). On the basis of this similarity, these proteins are thought to have similar functions. However, phylogenetic analyses of DRG1 and DRG2 have indicated that DRG1 and DRG2 belong to separate evolutionary branches; however, this finding has not been studied further (Etheridge et al. 1999; Li & Trueb 2000). Therefore, we hypothesized that DRG1 and DRG2 have distinct functions. We had previously proposed that transcription and/or stability of drg1 and drg2 mRNAs are regulated differently (Ishikawa et al. 2003). However, there has been no convincing evidence to differentiate the functions of DRG1 and DRG2.

Here we report the identification of two DRG family regulatory proteins (DFRPs) that have distinct binding specificities for DRG1 and DRG2. The association of one identified protein, DFRP1, with DRG1 is essential for maintenance of normal levels of expression of DRG1 in vivo, whereas DFRP1 does not bind DRG2. These findings suggest that DRG1 and DRG2 may be governed by distinct regulations in vivo.


   Results
Top
 Abstract
 Introduction
 Results
Discussion
 Experimental procedures
 References
 
Identification of the DRG family binding proteins

While searching databases of comprehensive analyses of protein complexes in yeast (Uetz et al. 2000; Ito et al. 2001; Ho et al. 2002), we noticed that Saccharomyces cerevisiae DRG1 and DRG2 interact physically with GIR2 (genetically interacting with ribosomal genes 2). GIR2 is highly conserved among eucaryotes (Homo sapiens GIR2 and S. cerevisiae GIR2 share 42% similarity at the amino acid (aa) level), indicating that it is involved in fundamental pathway. GIR2 has a characteristic RWD domain (Fig. 1A) (also called GI domain; Kubota et al. 2000; Doerks et al. 2002) that shows significant three-dimensional homology to ubiquitin conjugating-enzyme (E2) and E2 variant (UEV) (Nameki et al. 2004), suggesting that the DRG family may be regulated by poly-ubiquitination. To confirm that mammalian DRG family proteins also bind the mammalian homolog of GIR2, we amplified mouse DRG1, DRG2, and GIR2 cDNAs by PCR and constructed expression vectors. 293T cells were co-transfected with expression vectors for FLAG-tagged DRG1 or DRG2 together with Myc-tagged GIR2, and the cell extracts were subjected to immunoprecipitation assays with anti-FLAG antibody. The precipitates were analyzed by Western blotting. Myc-GIR2 was co-precipitated with FLAG-DRG2, whereas less Myc-GIR2 was co-precipitated with FLAG-DRG1 (Fig. 1B, left panels). These findings suggested that mouse GIR2 preferentially binds DRG2 rather than DRG1. Previously reported phylogenetic data showing that DRG1 and DRG2 are separated evolutionarily caused us to hypothesize that DRG1 might have a unique binding partner. Therefore, we screened for proteins structurally related to GIR2 in silico. A BLAST search with the full-length mouse GIR2 protein sequence as the query yielded a protein known as H. sapiens likely ortholog of mouse immediate early response erythropoietin 4 (LEREPO4). Both GIR2 and LEREPO4 have high lysine (K), glutamic acid (E), and aspartic acid (D) contents (GIR2, K, 8.2%; E, 16.2%; D, 9.1%; LEREPO4, K, 14.1%; E, 12.7%; D, 9.2%) and a highly homologous region that consists of approximately 60 amino acids, defined by alignment of multiple sequences from mouse, fly, and yeast (Fig. 1A, upper section, hatched boxes). However, sequences outside this region are not structurally similar. LEREPO4 has two unique CCCH-type zinc (Zn) fingers, which we termed ZnF-1 and ZnF-2 here, and a leucine-rich NES sequence that fits the widely accepted NES consensus [L-X2-3-(F, I, L, V, M)-X2-3-L-X-(L, I)] (Bogerd et al. 1996) at the C-terminus. LEREPO4 is also highly conserved among eucaryotes (H. sapiens LEREPO4 and S. cerevisiae LEREPO4 share 55% similarity at the aa level). To study the interactions between LEREPO4 and DRG family proteins, 293T cells were co-transfected with expression vector for FLAG-DRG1, or DRG2 together with vector expressing Myc-tagged LEREPO4, and the cell extracts were subjected to immunoprecipitation with anti-FLAG antibody followed by Western blotting. Myc-LEREPO4 was co-precipitated with FLAG-DRG1 but not with FLAG-DRG2 (Fig. 1B, right panels), indicating that LEREPO4 interacts selectively with DRG1. On the basis of the unique binding specificities of LEREPO4 and GIR2 for DRG1 and DRG2, respectively, we renamed LEREPO4 DRG family regulatory protein (DFRP) 1 and GIR2 DFRP2. In addition, the region that is highly homologous between DFRP1 and DFRP2 was termed DFRP domain (Fig. 1A, upper section, hatched boxes).



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  		Figure 1  Identification of DFRPs. (A) Domain architecture of mouse DFRP2 (GIR2) and DFRP1 (LEREPO4) and partial sequence alignments. DFRP1 and DFRP2 share a highly homologous region that we termed the ‘DFRP domain’ (hatched boxes). Multiple alignments of peptide sequences of M. musculus (Mm), D. melanogaster (Dm), and S. cerevisiae (Sc) DFRP2 and DFRP1. NCBI accession numbers: XP_125585 (DFRP2_Mm), NP_651227 [GenBank] (DFRP2_Dm), NP_010436 [GenBank] (DFRP2_Sc), NP_081210 [GenBank] (DFRP1_Mm), NP_610401 [GenBank] (DFRP1_Dm), and NP_014734 [GenBank] (DFRP1_Sc). Other domains are described in the text. (B) Interaction of DFRP2 (GIR2) or DFRP1 (LEREPO4) with the DRG proteins. Extracts of 293T cells co-transfected with expression vectors for FLAG-tagged mouse DRG1 or DRG2 or empty control and Myc-tagged mouse DFRP2 or DFRP1 were incubated with anti-FLAG antibody. The immunoprecipitated (IP) complexes or cell lysates on Western blots were then probed with anti-Myc antibody to detect the DFRPs. The same membranes were stripped and reprobed with anti-FLAG antibody to detect the DRG proteins. (C) In vivo binding assay. Protein extracts from HeLa S3 cells were incubated with antibodies against DFRP1, DRG1, and control IgG. The immunoprecipitated complexes were then probed with anti-DRG1 antibody. The same membrane was stripped and reprobed repeatedly with anti-DFRP1 and anti-DRG2 antibodies. (D) Immunofluorescence analysis of the subcellular localization of DFRP1 and DRG1. HeLa S3 cells were stained with polyclonal antibodies against DFRP1 and DRG1.

 
In vivo physical interaction between DFRP1 and DRG1

To confirm the specific interaction of DRG1 and DFRP1 in vivo, we generated polyclonal antibodies that specifically recognize endogenous DFRP1, DRG1, or DRG2. To remove cross-reactive antibodies that recognized both DRG1 and DRG2, anti-DRG1 sera and anti-DRG2 sera were precleared by adsorption to recombinant DRG2 and DRG1, respectively, before final affinity purification. The purified antibodies against DRG1 and DRG2 were confirmed to be highly specific without recognizable cross-reactivity (data not shown). We then performed immunoprecipitation experiments using extracts from HeLa S3 cells to study whether endogenous DFRP1 associates with DRG1. DRG1 was detected by Western blot analysis with anti-DRG1 antibody following immunoprecipitation with anti-DFRP1 antibody, whereas DRG2 was not detected with anti-DRG2 antibody in the same immunoprecipitates (Fig. 1C, lane 2). In the reverse experiment, DFRP1 was identified following immunoprecipitation with anti-DRG1 antibody (Fig. 1C, lane 3). DRG1 and DFRP1 were not detected on Western blots following immunoprecipitation with unrelated rabbit control IgG (Fig. 1C, lane 4). These results indicate that endogenous DFRP1 associates specifically with DRG1 but not with DRG2.

To further confirm the association between endogenous DFRP1 and DRG1, the subcellular localization of DFRP1 and DRG1 was investigated by immunofluorescence microscopy. Because the antibodies for DFRP1 and DRG1 both originated from rabbits, double staining with these antibodies was not possible. Therefore, we stained HeLa S3 cells independently with anti-DFRP1 or anti-DRG1 antibody. In both cases, signals were detected throughout the cytoplasm of HeLa S3 cells in an identical pattern (Fig. 1D). Further analysis with subcellular localization vectors that encode proteins targeting various organelles (subcellular Localization Vectors, BD Biosciences Clontech) revealed that the distribution patterns of DFRP1 and DRG1 were not identical to those of endoplasmic reticulum, Golgi, and mitochondria (data not shown), suggesting that DFRP1 and DRG1 may be co-localized in cytosol. These data strongly support the idea that DFRP1 and DRG1 form a complex in vivo.

Requirement of DFRP domain for the interaction of DFRPs with DRG proteins

Next we determined which region of DFRPs is required for the interaction with the DRG family. We generated several deletion mutants of DFRP2 and DFRP1 (Fig. 2A). Deletion of 198–243 C-terminal residues and 1–141 N-terminal residues of DFRP2 (termed {Delta}C2 and {Delta}N1, respectively) did not affect binding to DRG2 (Fig. 2B, lanes 1, 3, 4). However, further truncated mutants ({Delta}C1 and {Delta}N2) lacking the most part of the DFRP domain did not bind DRG2 (Fig. 2B, lanes 2 and 5). Therefore, it appears that the DFRP domain of DFRP2 is essential for the interaction with DRG2. Deletion of the N-terminal and C-terminal halves of DFRP1 ({Delta}N and {Delta}C, respectively) did not significantly affect binding to DRG1 (Fig. 2C, lanes 1–3). This overlapping region is part of the DFRP domain and is highly conserved among DFRP1s in eucaryotes (Fig. 1A). We then constructed an interstitial deletion mutant of DFRP1 ({Delta}D1) that lacks the highly conserved region. This deletion completely abolished the ability to associate with DRG1 (Fig. 2C, lane 4). Therefore, a 25-aa peptide of the DFRP domain is essential for the interaction with DRG1. These results suggest that the DFRP domain is deeply involved in the interaction with the DRG family. Given the highly similar sequences of the DFRP domain in DFRP1 and DFRP2, it is possible that subtle differences in the structures of the DFRP domains of DFRP1 and DFRP2 are responsible for their different binding specificities.



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  		Figure 2  Requirement of the DFRP domain for association of DFRPs with DRG proteins. (A) Structural maps of DFRP2 (upper section) and DFRP1 mutants (lower section). Hatched boxes indicate DFRP domain. Gray solid bar corresponds to a highly conserved region of DFRP1s in eucaryotes as depicted in Figure 1A. (B) Determination of the essential region in DFRP2 for interaction with DRG2. Extracts from 293T cells co-transfected with expression vectors for GST-fusion DFRP2 fl or its mutants and FLAG-tagged DRG2 were incubated with glutathione sepharose beads. The pull-down precipitates or the cell lysates were then probed with anti-FLAG antibody to detect FLAG-DRG2. The membrane was stripped and reprobed with anti-GST antibody to detect GST-DFRP2 fl and its mutants. (C) Determination of the essential region in DFRP1 for interaction with DRG1. Extracts from 293T cells co-transfected with expression vectors for FLAG-tagged DFRP1 fl or its mutants and Myc-tagged DRG1 were incubated with anti-FLAG antibody. The immunoprecipitates (IP) or the cell lysates were then probed with anti-Myc antibody to detect Myc-DRG1. The membrane was stripped and reprobed with anti-FLAG antibody to detect FLAG-DFRP1 fl and its mutants.

 
Regulation of expression of DRG proteins by DFRPs

In our experiments, we were unable to produce over-expression of DRG1 or DRG2 alone by transient transfection of expression vector for DRG1 or DRG2 (Fig. 3A, lanes 5 and 6). This phenomenon was previously reported for DRG1 (Sazuka et al. 1992b; Mahajan et al. 1996). However, when we co-expressed DRG1 with DFRP1 or DFRP2, expression of DRG1 increased dramatically (Fig. 3A, lanes 1 and 3). Expression of DRG2 was also increased when it was coexpressed with DFRP2 but not significantly with DFRP1 (Fig. 3A, lanes 2 and 4). We hypothesized that exogenous or over-expressed DRG proteins may be targeted for degradation by an unknown mechanism and that DFRPs may inhibit such degradation through physical association with DRGs. Because DFRP1 has two Zn-finger domains, which could interact with ubiquitin, and because DFRP2 has an RWD domain, which is structurally related to ubiquitin-conjugating enzymes (E2s), we hypothesized that DFRPs protect DRGs from degradation by the ubiquitin/proteasome system. To test this possibility, 293T cells were co-transfected with expression vectors for HA-ubiquitin, FLAG-DRG1, or DRG2 and incubated with or without the 26S proteasome inhibitor MG132 for 3 h. Cell extracts were immunoprecipitated with anti-FLAG antibody followed by Western blotting with anti-HA antibody (Fig. 3B, IP blots; note that protein samples for IP blots were normalized for amounts of DRG proteins). Accumulation of poly-ubiquitin conjugates in DRG1 or DRG2 immunocomplexes were detected upon addition of MG132 (Fig. 3B, IP blots, top panels, lanes 2 and 8), whereas such accumulations were not observed in the absence of MG132 (lanes 1 and 7). This suggests that DRG proteins may be constitutively degraded through a ubiquitination-dependent mechanism. On addition of MG132, no accumulation of native-size of DRG proteins was observed, suggesting that ubiquitination of DRG proteins may not be affected by the MG 132 treatment (Fig. 3B, Lysate blots, upper panels, lanes 1, 2 and 7, 8; note that protein samples for Lysate blots were normalized for cell number). Co-expression of Myc-DFRP1 with DRG1 or DRG2 leads to accumulation of DRG1 in a dose-dependent manner (Fig. 3B, Lysate blots, upper panels, lanes 2–4). Accumulation of DRG2 was not significantly observed (lanes 8–10). Co-expression of Myc-DFRP2 resulted in the accumulation of both DRG1 and DRG2 (lanes 2, 5, 6 and 8, 11, 12). Furthermore, increased DFRP1 expression reduced levels of poly-ubiquitin conjugates in DRG1 immunocomplexes (Fig. 3B, IP blots, top panels, lanes 2–4) but not significantly in DRG2 immunocomplexes (lanes 8–10). DFRP2 reduced poly-ubiquitination of both DRG1 and DRG2 immunocomplexes (lanes 2, 5, 6 and 8, 11, 12). In the same immunoprecipitated complexes, a specific interaction between DRGs and DFRPs was confirmed with an anti-Myc antibody (Fig. 3B, IP blots, bottom panels). These results suggest that in transient transfection experiments, DFRP1 specifically stabilizes DRG1, whereas DFRP2 stabilizes both DRG1 and DRG2 by protecting them from poly-ubiquitination followed by proteolysis.



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  		Figure 3  Regulation of expression of DRG proteins by DFRPs. (A) Increased expression of DRG proteins by coexpression with DFRPs. Extracts of 293T cells transfected with an expression vector for Myc-tagged DRG1 or DRG2 alone or together with an expression vector for FLAG-tagged DFRP1 or DFRP2 were analyzed by Western blotting with anti-Myc and anti-FLAG antibodies. (B) Inhibition of ubiquitination of DRG proteins by association of DFRPs. 293T cells transfected with the expression vectors encoding HA-Ubiquitin (Ub) (3 µg) and FLAG-DRG1 (left panels) or DRG2 (right panels) (3 µg), and together with Myc-DFRP1 or DFRP2 (1 or 5 µg) were treated with (+) or without (–) 10 µM of MG132 3 h before harvest. IP panels, cell lysates were incubated with anti-FLAG antibody, and immunoprecipitated complexes were probed with anti-HA antibody to detect ubiquitin conjugates. The same membrane was stripped and reprobed repeatedly with anti-FLAG and anti-Myc antibody to detect DRGs and DFRPs, respectively. Black arrows and white arrows indicate full-length Myc-DFRP1 and Myc-DFRP2, respectively. On these membranes, the amounts of FLAG-DRG1 and FLAG-DRG2 were equalized on all lanes. To do this, the loading amount of each sample was determined by preliminary assessment of the concentration on another blot with the same samples. Lysate panels, cell lysates used for immunoprecipitation reactions were analyzed by Western blotting with anti-Myc antibody to detect DFRPs. The same membrane was stripped and reprobed with anti-FLAG antibody to detect DRG1 and DRG2. On SDS-PAGE, we loaded extracts of equal numbers of cells.

 
Regulation of DRG1 protein expression by DFRP1 in vivo

To examine DFRP1-mediated regulation of DRG1 protein expression in vivo, we generated a DFRP1-deficient DT40 chicken B cell line. We first isolated a partial cDNA fragment of chicken dfrp1 by PCR with degenerate primers designed from the conserved portions of H. sapiens and Drosophila melanogaster DFRP1 peptide sequences. We used the sequence of the amplified cDNA to design primers to amplify a partial fragment of the chicken dfrp1 gene locus by long PCR. Subsequent sequence analysis of the isolated genomic DNA revealed that the locus contains several exons, one of which encodes full-length Zn-finger 1 domain (ZnF-1, illustrated in Fig. 1A). To target disruption of the ZnF-1 exon, we constructed targeting vectors (dfrp1Bsr and dfrp1HisD) in which a blasticidin- or a histidinol-resistance (Bsr or HisD) gene cassette was flanked by 5'- and 3'-genomic arms situated upstream or downstream of the exon, as illustrated in Fig. 4A. After wild-type DT40 cells were transfected with dfrp1Bsr, blasticidin-resistant clones were isolated. One of these heterozygous clones was then transfected with dfrp1HisD to delete the second allele. Both targeting events were confirmed by Southern blot analysis of genomic DNA with 5'-flanking probe (Fig. 4B). Additional proof of dfrp1 elimination was provided by Northern and Western blot analyses (Fig. 4C,D, respectively).



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  		Figure 4  Disruption of the dfrp1 gene in DT40 cells. (A) Structure of the partial chicken (Gallus gallus) dfrp1 gene locus and knockout constructs. Hatched boxes represent exons of the chicken dfrp1 gene. The left exon contains the ZnF-1 domain. To target this exon, blasticidin (Bsr)- and histidinol (HisD)-resistance genes were flanked by the upstream and downstream genomic arms. The resultant targeting vectors were termed dfrp1Bsr and dfrp1HidD, respectively. The locations of the ApaI (A) and BamHI (B) restriction sites, used in the Southern blot analysis of possible recombinant clones are shown beside the maps. (B) Southern blot analysis of ApaI-BamHI-double digested genomic DNA prepared from DT40 wild-type (+/+) and dfrp1+/– or dfrp1–/– cells. (C) Northern blot analysis of dfrp1 mRNA expression in DT40 wild-type (+/+) and dfrp1–/– cells. Ethidium bromide (EtBr)-stained rRNAs are shown as loading controls. (D) Western blot analysis of DT40 wild-type (+/+) and dfrp1–/– cells. Tubulin was stained as an internal loading control.

 
Targeted deletion of dfrp1 (dfrp1–/–) resulted in a dramatic reduction in DRG1 expression, whereas tubulin control expression was not affected (Fig. 5A,a,d); this indicates that DFRP1 is required for normal expression of DRG1 in vivo. To verify that this phenotype was due solely to the deficiency of dfrp1, we introduced vectors expressing murine full-length (mfl) or mutant (m{Delta}D1) DFRP1, which lacks aa essential for interaction with DRG1, into dfrp1–/– cells. We established chicken dfrp1-deficient cells that stably expressed DFRP1fl (dfrp1–/–mfl) or DFRP1{Delta}D1 (dfrp1–/–m{Delta}D1). In the dfrp1–/–mfl cells, expression of DRG1 was restored (Fig. 5A,a, compare lanes 1–3), indicating that the reduction of DRG in dfrp1–/– cells was due to lack of DFRP1. However, in dfrp1–/–m{Delta}D1 cells, expression of DRG1 was not restored (lane 4). This result strongly suggests that binding of DFRP1 to DRG1 is essential to maintain normal levels of DRG1. To study whether the reduction of DRG1 expression occurs at the transcriptional or post-transcriptional level, drg1 mRNA levels were analyzed by Northern blot analysis. Levels of drg1 mRNA did not differ significantly in the presence or absence of DFRP1 or its mutants (Fig. 5A,e). Therefore, the observed changes in DRG1 expression in the present experiment were not due to changes in drg1 expression. This suggests that regulation of DRG1 expression by DFRP1 occurs at the post-transcriptional level. In contrast, DRG2 levels were similar across all cell types (Fig. 5A,b). Northern blot analysis revealed that the levels of drg2 mRNA in dfrp1–/– and dfrp1–/–m{Delta}D1 cells are higher than levels in wild-type and dfrp1–/–mfl cells (Fig. 5A,f), although this phenomenon is not clear at present. These results indicate that DFRP1 specifically up-regulates levels of DRG1 in vivo.



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  		Figure 5  Regulation of DRG1 expression by DFRP1 in vivo. (A) Western blot (upper panels) and Northern blot (lower panels) analysis of DT40 wild-type, dfrp1–/–, dfrp1–/–mfl, and dfrp1–/–m{Delta}D1 cells. Whole-cell extracts were separated by SDS-PAGE and then transferred to a membrane that was immunoblotted repeatedly with antibodies against DRG1, DRG2, DFRP1, and tubulin (for internal loading control). Total cell numbers per lane were equalized (2.5 x 105 cells/lane). For Northern blot analysis, cells were collected from the same culture dish used for Western blot analysis. Total RNA was isolated, separated through a 1.2% formaldehyde denaturing gel, transferred to a nylon membrane, and probed with radiolabelled chicken drg1, drg2, or gapdh (for internal control) partial cDNA probes. The same membrane was used repeatedly. Band intensity was quantified on a Fujifilm BAS2000 bio-imaging autoanalyser. Results are displayed as the ratio of expression in dfrp1–/–, dfrp1–/–mfl, or dfrp1–/–m{Delta}D1 cells to that in wild-type cells. (B) Immunoprecipitation analysis for association between DFRP1 and DRG1. Numbers of DT40 wild-type, dfrp1–/–, dfrp1–/–mfl, and dfrp1–/–m{Delta}D1 cells were adjusted to 1 : 7 : 1 : 7, respectively, to equalize the total amount of DRG1 protein in each extract. Cell extracts were incubated with anti-DFRP1 antibody, and the immunoprecipitated complexes and cell lysates were then probed with either anti-DRG1 or anti-DFRP1 antibody on Western blot analysis.

 
To address the question of whether the DFRP1-mediated regulation of expression of DRG1 is caused by the physical interaction of DFRP1 with DRG1, we carried out in vivo immunoprecipitation assays. Immunoprecipitation with anti-DFRP1 antibody followed by immunoblotting with anti-DRG1 antibody revealed that endogenous DFRP1 is associated with DRG1 in DT40 wild-type cells (Fig. 5B, lane 1). Stably expressed mDFRP1fl was also associated with DRG1, whereas mDFRP1{Delta}D1 was not bound to DRG1, even though the level of mDFRP1{Delta}D1 expression was sufficient (Fig. 5B, lanes 3, 4). In the reverse experiment, we used anti-DRG1 antibody for immunoprecipitation and anti-DFRP1 antibody for immunoblotting, and the result led to the identical conclusion (data not shown). These results strongly suggest that normal expression of DRG1 requires physical association with DFRP1 in vivo.

Spatial and temporal co-expression of drg1 and dfrp1 in X. laevis

drg1 was first identified as a gene expressed predominantly during early development of the mouse central nervous system, and we previously reported a comparative analysis of drg1 and drg2 expression in X. laevis embryos and adult tissues (Ishikawa et al. 2003). Whole mount in situ hybridization and Northern blotting revealed different expression patterns for drg1 and drg2. For example, only drg2 expression is detected in stage 22 of pronephric anlage. To determine whether DFRP1 acts as a regulator of DRG1 universally, we used X. laevis to compare spatial and temporal expression of drg1 and dfrp1 during embryogenesis and in various adult tissues. We first cloned the Xenopus dfrp1 cDNA as described in experimental procedures. Whole mount in situ hybridization of Xenopus embryos revealed that the expression patterns of dfrp1 were quite similar to those of drg1 (Fig. 6A,a–j). At stage 22, both genes were expressed in blood islands, somites, developing eyes, trunk neural crest, mandibular crest segment, hyoid crest segment, and branchial crest segment (Fig. 6A,a–f). At this stage, neither dfrp1 nor drg1 was expressed at the region of pronephric anlage, whereas drg2 mRNA levels were high at pronephric anlage (Ishikawa et al. 2003), suggesting that the transcription and/or stability of dfrp1 mRNA may be regulated in a manner more similar to that of drg1 than to that of drg2. At stage 32, the expression patterns of drg1 and dfrp1 were almost identical: both genes were expressed in otic vesicle, pronephros, forebrain, midbrain, hindbrain, branchial arch, eyes, lens, spinal cord, and notochord (Fig. 6A,g–j). In adult tissues, dfrp1 was expressed strongly in ovary; moderately in brain, kidney, spleen, testis, intestine, and colon; and scarcely in heart, lung, liver, stomach, and skeletal muscle (Fig. 6B). This expression pattern is more similar to that of drg1 than that of drg2, which is expressed at moderate levels in heart, lung, and liver (Ishikawa et al. 2003). We also examined the temporal expression of dfrp1 during early stages of development (Fig. 6C). Expression of dfrp1 was induced weakly in late gastrula (stages 13–14) and strongly from late neurula (stages 20–22) to tadpole (stages 40–41). This pattern is similar to that of drg1 (Ishikawa et al. 2003). These spatial and temporal expression similarities between dfrp1 and drg1 in a multicellular organism support the idea that DFRP1 may cooperate with DRG1 in various types of cells.



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  		Figure 6  Expression analysis of dfrp1 and drg1 in X. laevis.(A) Spatial expression of dfrp1 and drg1 transcripts during Xenopus embryonic development. (a,b) Ventral views, anterior left; (c,d) Dorsal views, anterior left; (e–j) Lateral views, anterior left; (g' and h') Higher magnification images of the anterior part of the embryo depicted in g and h, respectively; (i, j) Clearing of embryo by benzyl alcohol:benzyl benzoate (2 : 1). Abbreviations: ba, branchial arch; bcs, branchial crest segment; bi, blood islands; de, developing eyes; e, eyes; fb, forebrain; hb, hindbrain; hcs, hyoid crest segment; le, lens; mb, midbrain; mcs, mandibular crest segment; nc, notochord; ov, otic vesicle; pr, pronephros; sc, spinal cord; sm, somite; tnc, trunk neural crest. (B) Tissue-specific expression of dfrp1 mRNAs in adult Xenopus. Total RNAs were isolated from the indicated adult tissues for Northern blotting. (C) Temporal expression of dfrp1 transcripts during X. laevis embryonic development. Total RNAs were isolated from Xenopus embryos at the indicated stages for Northern blotting. In (B,C), the same membrane that was used in our previous paper (Ishikawa et al. 2003) was reprobed. Therefore, quality and amount of RNA loaded were confirmed.

 

   Discussion
Top
 Abstract
 Introduction
 Results
 Discussion
Experimental procedures
 References
 
In the present study, we identified a novel family of two proteins, DFRP1 and DFRP2, that are highly conserved in eucaryotes and specifically bind to and regulate the expression of target DRG proteins through blockade of poly-ubiquitination that could lead to degradation (Figs 1B and 3). One interesting feature of the DFRPs is the presence of a unique domain that we have termed the DFRP domain and that shows a high sequence identity among the DFRPs (Fig. 1A). DFRP domain is essential for binding to the DRG family (Fig. 2). Therefore, binding of DFRPs to members of the DRG family and maintenance of the proper expression of DRG proteins may be sufficiently important in eukaryotic organisms to necessitate evolutionary conservation. It is noteworthy that DRG1 protein is also stabilized when SCL (TAL-1), a transcription factor for hematopoietic development that can bind DRG1, is transiently coexpressed (Mahajan et al. 1996). It appears that degradation occurs when DRG proteins are released from stable protein complexes. Such degradation of free DRG molecules might be mediated by poly-ubiquitination through the concerted actions of ubiquitin activase (E1), conjugase (E2), and ligase (E3). Further studies are needed to determine which catalytic enzymes ‘monitor’ free DRG family proteins and target them for degradation. Specific domains in DFRPs may be linked to these ubiquitin-related enzymes. DFRP2 has an RWD domain, which is frequently found in proteins that contain other domains, such as RING finger, IBR (In Between Ring fingers), UBA, UBCc, and WD repeat domains, characteristic of proteins involved in degradation through the ubiquitin-proteasome pathway. Furthermore, two consecutive repeats of the CCCH-type Zn finger in DFRP1 are similar to the C3HC4-type and C3H2C3-type RING fingers. We believe that these two Zn-finger repeats may be a RING finger variant. Moreover, a single Zn finger can associate with ubiquitin. For example, the novel zinc finger (NZF) domain is found in NPL4 that is involved in ER-associated degradation (ERAD) (Wang et al. 2003). The polyubiquitin-associated zinc finger (PAZ) domain is found in microtubule-associated HDAC6 (Hook et al. 2002). However, the effects of DFRP1 and DFRP2 are the reverse of what would be expected; they prevent, not promote, degradation of DRGs. One possible explanation of this inconsistency is that the DFRP domain is a pseudo-domain that interacts with DRGs like its real counterpart but cannot target the protein for degradation. Another possibility is that the RWD or two Zn-finger motifs in DFRPs have deubiquitination activity. The true function of these domains remains unclear and needs further study.

On the basis of previous studies, DRG proteins may play critical roles on cell growth. drg1, drg2, and dfrp1 transcripts are expressed at high levels in growing Xenopus embryos (Fig. 6; Ishikawa et al. 2003). In Pisum sativum and Arabidopsis thaliana, drg2 mRNA accumulates preferentially in growing tissues (Devitt et al. 1999; Etheridge et al. 1999). Furthermore, LEREPO4, which we renamed DFRP1, was originally identified as a gene transcribed immediately in response to erythropoietin (Epo) signalling via a C-terminal-truncated Epo receptor in erythroleukaemic SKT6 cells (Gregory et al. 2000). Epo receptor relays key signals for growth (Shikama et al. 1996). If stabilization is necessary for the cellular function of DRG1, DRG1 may start its function upon induction of DFRP1 by such growth signals. However, aberrant expression of the DRG family leads to cell transformation or cell cycle arrest (Mahajan et al. 1996; Ko et al. 2004; Song et al. 2004). On the basis of our data, over-expressed DRG proteins that may escape ubiquitin-mediated degradation could function incorrectly. Gene disruption studies as well as further analysis of the unique regulations by DFRP1 and DFRP2 may provide the best clues as to the functions of the DRG family proteins.


   Experimental procedures
Top
 Abstract
 Introduction
 Results
 Discussion
 Experimental procedures
References
 
Reagents and plasmids

Blasticidin, histidinol, and puromycin were purchased from Sigma. MG132 was purchased from the Peptide Institute. FLAG-tagged, Myc-tagged and GST-fusion expression vectors were constructed by insertion of cDNA fragments into pME-FLAG, pME-Myc or pME-GST modified from the SR{alpha} promoter-driven expression vector pME18S (Shiio et al. 1992). cDNA fragments of mouse DFRP1fl, DFRP1{Delta}N (aa 231–426), DFRP1{Delta}C (aa 1–263), DFRP2, DRG1, and DRG2 were amplified by PCR from a mouse neural tube cDNA library fused to pAD-GAL4-2.1 (Stratagene). DFRP2 cDNA was digested with NcoI, filled in with Klenow and used to generate {Delta}C1 and {Delta}N1 mutants of DFRP2. For generating {Delta}C2 and {Delta}N2 mutants of DFRP2, EcoRV site was utilized. DFRP1{Delta}D1 (deleted aa 236–260) was generated by inserting a BglII site into the appropriate position between two BglII sites of DFRP1 cDNA by the Kunkel method and inserting the BglII-digested larger fragment into BglII-digested pME-FLAG-DFRP1fl vector. HA-tagged ubiquitin was cloned into pcDNA3.1 vector (Invitrogen). The drug-resistance gene cassette, pA-puro vector for the construction of DT40 stable cell lines, and DT40 genomic library were gifts from Dr Kurosaki (RIKEN, Japan).

Antibodies

The cDNAs encoding mouse DFRP1 (aa 1–270, named {Delta}C2), DFRP2, DRG1, and DRG2 were subcloned into pGEX-4T-1 (Amersham Biosciences) and pMAL-c (New England BioLabs) expression vectors. The vectors were transformed into Escherichia coli (BL21). The expressed GST-fusion and MBP-fusion proteins (GST-DFRP1{Delta}C2, GST-DFRP2, GST-DRG1, GST-DRG2, MBP-DRG1, and MBP-DRG2) were isolated from bacterial lysates with glutathione sepharose 4B beads (Amersham Biosciences) and amylose resin (New England BioLabs), respectively, according to the manufacturer's instructions. To generate antisera, rabbits were immunized subcutaneously at 2-week intervals with GST-DFRP1{Delta}C2, GST-DFRP2, MBP-DRG1, or GST-DRG2 emulsified initially in 50% Freund's complete adjuvant and subsequently in 50% Freund's incomplete adjuvant (Sigma). DFRP1{Delta}C2 antiserum was affinity purified with MAbTrapTM.GII columns (Amersham Biosciences). Antiserum against GST-DFRP2 was adsorped to GST protein that were immobilized to N-hydroxysuccinimide (NHS) HiTrap columns (Amersham Pharmacia Biotech) to obtain anti-GST antibodies. Antisera against MBP-DRG1 and GST-DRG2 were affinity purified by final adsorption to GST-DRG1 and MBP-DRG2 columns, respectively. To reduce cross-reactivity, the antisera against DRG1 and DRG2 were precleared by adsorption to MBP-DRG2 and GST-DRG1 columns, respectively, before proceeding to the final columns. Anti-c-Myc (A-14) and anti-HA (F-7) antibodies were purchased from Santa Cruz Biotechnology; anti-FLAG (M2) was from Sigma. Anti-tubulin (Ab-1) was from Oncogene.

Immunoprecipitation assay

For immunoprecipitation, cultured cells were lysed in TNE buffer (10 mM Tris-HCl, pH 7.8, 1% Nonidet P-40, 150 mM NaCl, 1 mM EDTA). The cell lysates were centrifuged and the supernatants were precleared by incubation with protein-G Sepharose beads (Amersham Biosciences). Cleared lysates were incubated with appropriate antibodies for 1 h with protein-G Sepharose beads. Beads were collected by centrifugation and washed three times with TNE buffer. Captured proteins were eluted from the beads by boiling in SDS sample buffer and analyzed by SDS-PAGE, followed by Western blotting. For analysis of ubiquitin complexes, vector-transfected cells were allowed to grow for 2 days and were treated or not treated with 10 µM of proteasome inhibitor MG132 3 h before lysis in TNE buffer.

Immunofluorescence analysis

HeLa S3 cells were cultured on glass coverslips and allowed to grow for 2 days. Attached cells were washed in PBS, fixed in methanol-acetone (1 : 1) for 10 min, dried, and blocked in 2% BSA. The fixed cells were incubated with antibody against DFRP1 or DRG1 for 1 h. At the end of incubation, cells were washed in PBS containing 0.2% Tween20 and stained with an Alexa 488-conjugated anti-rabbit secondary antibody (Molecular Probes) for 1 h. Slides were examined with a laser scanning confocal microscope (Radiance 2000, Bio-Rad). Z series consisting of three images were collected through the depth of the cells with an iris setting of 2.0 and a step size of 0.5 µm. Images were projected with the maximum pixel method (LaserSharp 2000 software, Bio-Rad).

DT40 cells

DT40 cells were grown in RPMI medium (JRH Biosciences) supplemented with 10% fetal calf serum (Sigma), 1% chicken serum (Sigma), penicillin, streptomycin, and ß-mercaptoethanol. A fragment of the chicken DFRP1 cDNA was amplified from DT40 cDNA with a set of degenerate primers (5'-GCGAATTCATGCCNCCNAARAARC-3' and 5'-GCCTCGAGYTTYTCYTCYTTYTTYTTRTC-3'). Full-length DFRP1 cDNA (DDBJ/EMBL/GENBANK Accession no. AB185935) was isolated by screening approximately 5 x 106 plaques of a DT40 cDNA library ({lambda}Zap) with the partial cDNA fragment as a probe. A genomic DNA fragment containing part of the dfrp1 gene locus was isolated by long PCR with LA Taq and forward primer 5'-AGCAAGAAGGCGGACCAGAA-3' and reverse primer 5'-GAGGAAGAGCATGGCGATAC-3'. To construct the targeting vectors dfrp1Bsr and dfrp1HisD for disruption of the chicken dfrp1 gene, approximately 2.5 kb of genomic DNA containing an exon encoding the ZnF-1 domain was replaced by a blasticidin- or histidinol-resistance gene in reverse orientation to the transcription of dfrp1. Stable transfectants following electroporation (Gene Pulser II, Bio-Rad, 550 V, 25 µF) of pA-puro vectors inserted with mouse DFRP1 fl or {Delta}D1 were selected by growth in puromycin. For Northern blot analysis, total RNA was hybridized to the chicken dfrp1 cDNA probe (nt +1 to +1290), a chicken drg1 partial cDNA probe (562 bp, amplified by PCR with degenerate primers 5'-GGCAGCCTAYGAATTYAC-3' and 5'-AAARTTCCAGCGGTGATG-3', DDBJ/EMBL/GENBANK accession no. AB186130), or the chicken drg2 partial cDNA probe (564 bp, amplified by PCR with primers 5'-GCGAATTCTGCATCTTATGAGTTCAC-3' and 5'-GCCTCGAGCCAGGTTCAATTTCATG-3').

Expression analysis in Xenopus laevis

To isolate Xenopus dfrp1 cDNA, we first amplified a partial cDNA fragment encoding a peptide highly homologous to the N-terminal portion of mouse DFRP1 by PCR from Xenopus liver cDNA pool using forward primer 5'-GCGGATCCATGCCGCCTAAGAAAG-3' and reverse primer 5'-GCCTCGAGGCGTCATCTGCTTCTT-3'. We used this fragment to probe approximately 7 x 105 plaques of a Xenopus laevis embryo (stage 24–32) cDNA library in {lambda}ZipLox (Gibco BRL) at high stringency (0.2 x SSC, 0.1% SDS, 50 °C). The complete sequence of the isolated Xenopus dfrp1 cDNA was submitted to DDBJ/EMBL/GENBANK (Accession no. AB185934). The procedures of whole mount in situ hybridization and Northern blot analysis were previously described (Ishikawa et al. 2003). We used cDNA template covering nucleotides –68 to +1755 for Xenopus dfrp1 probe.

Computational analysis

Multiple sequences were aligned by ClustalX software (ver. 1.81). Manual modification was added to the aligned results for sophistication. The aligned data were shaded by BOXSHADE software (ver. 3.31). Characteristic domains and positions (Fig. 1A) were determined by a SMART database search and published descriptions (Bogerd et al. 1996).


   Acknowledgements
 
We thank Dr N. Sato for technical support, Dr T. Kurosaki for the DT40 genomic library, and Dr K. Tanaka for HA-tagged ubiquitin vector. This work was supported by a grant-in-aid for Scientific Research on Priority Areas and by Special Coordination Funds for Promoting Science and Technology from the Ministry of Education, Culture, Sports, Science and Technology of Japan.


   Footnotes
 
Communicated by: Shunsuke Ishii

* Correspondence: E-mail: jun-i{at}ims.u-tokyo.ac.jp


   References
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 Introduction
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 Discussion
 Experimental procedures
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Received: 20 September 2004
Accepted: 17 November 2004





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