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1 Division of Life Science and Biotechnology, Department of Ecological Engineering, Toyohashi University of Technology, Toyohashi, Aichi 441-8580, Japan
2 Cellular Physiology Laboratory, RIKEN, Wako, Saitama 351-0198, Japan
3 Graduate School of Frontier Biosciences, Osaka University, Suita, Osaka 565-0871, Japan
4 SORST, Japan Science and Technology Agency, Suita, Osaka 565-0871, Japan
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Abstract |
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 Top Abstract IntroductionResultsDiscussionExperimental proceduresReferences |
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Introduction |
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TopAbstractIntroductionResultsDiscussionExperimental proceduresReferences |
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Results |
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TopAbstractIntroductionResultsDiscussionExperimental proceduresReferences |
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We previously identified drh-3(D2005.5) as a candidate that could be implicated in maintenance of genome integrity during systematic RNAi-mediated screens of C. elegans helicase-related genes (unpublished data). Progenies from the nematode C. elegans, where the candidate gene was treated with RNAi, exhibited increased sensitivity to X-ray irradiation. The protein motif search revealed that the drh-3 gene encodes a novel RNA helicase-like protein with a typical DEXDc motif (smart00487) that is conserved in the DEAD-like helicases superfamily. The BLASTP search, using DRH-3 as a query, hit several RNA helicases, including Dicer proteins that play a crucial role in RNAi in many species including C. elegans (Knight & Bass 2001) (data not shown). The amino acid sequences corresponding to the representative proteins from identified DRH-3-like proteins were aligned and clustered into subgroups based on their sequence similarities. A part of the resultant alignment of 11 representative proteins shown in Fig. 1A indicates that DRH-3 contains several typical helicase motifs, such as motifs I and II (Walker motif A and B, respectively) for ATP binding and hydrolysis (Walker et al. 1982). Interestingly, the alignment shows that nematode DRH-2 protein does not contain motif I (Walker motif A) (Fig. 1A). The phylogenic tree from the clustering of 32 sequences, shown in Fig. 1B, indicates that the DRH-3 protein belongs to the subgroup of Dicer-related helicase (DRH) proteins that include the C. elegans DRH-1 and DRH-2 proteins. Both DRH-1 and DRH-2 are required for RNAi in the nematode (Tabara et al. 2002). Recently, DRH-3 was identified as the third member of this group (Duchaine et al. 2006). Members belonging to another subgroup, denoted by IFIH1/DDX58, are very similar to DRH proteins. The subgroup contains mammalian RNA helicases DDX58 [also known as RIG-1 (retinoic acid-inducible gene 1)] (Yoneyama et al. 2004), IFIH1 (interferon induced with helicase C domain 1, also known as MDA-5) (Kang et al. 2002) and LGP2 (Yoneyama et al. 2005), which have all been implicated in signal transduction in the antiviral response. The DRH proteins also share sequence similarities to the members of two other subgroups, denoted DICER and Hef/Mph1p/FANCM. The proteins in the DICER subgroup act exclusively in RNAi pathways in many eukaryotes (Jaronczyk et al. 2005), and several proteins in the Hef/Mph1p/FANCM subgroup appeared to play roles in DNA repair processes including the FA pathway in mammals (Kennedy & D’Andrea 2005).
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To detect the transcript from drh-3 gene in hermaphrodites, we performed reverse transcription PCR (RT-PCR) analyses using RNA isolated from nematode. Total RNA was prepared from the animals synchronized at different developmental stages [i.e. the first-to-second (L1–L2) stage larvae, the L3–L4 stage larvae, young adult animals and adult animals]. Semi-quantitative RT-PCR was performed using specific primers for drh-3 and for an internal control gene, ama-1, which encodes the large subunit of RNA polymerase II. The drh-3 and ama-1 transcripts in the reactions were detected as shown in Fig. 2. The level of ama-1 transcript in RNA isolated from animals at each developmental stage was similar with each other. In contrary, levels of the drh-3 transcripts in animals at adult stages were approximately threefold higher than that in larval nematodes (Fig. 2). These data may suggest that drh-3 plays a role in biological processes required for adults rather than for larvae.
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To investigate the role of the DRH-3 protein in C. elegans, drh-3 gene expression was suppressed by feeding worms Esherichia coli containing plasmid vectors designed for the production of double-stranded RNA (dsRNA) by the bidirectional transcription of inserts (Timmons et al. 2001). Larvae at the L1 stage were fed bacteria producing dsRNA specific to drh-3 or control bacteria transformed by the vector alone (i.e. mock treatment) and grown to adulthood. The levels of drh-3 transcripts after feeding RNAi were examined with RT-PCR on total RNA from these RNAi-treated animals using specific primers for drh-3 and for a control gene, ama-1. Figure 3A shows that drh-3 mRNA was depleted in drh-3(RNAi) worms (lane 4) but not in mock-treated animals (lane 3), while the control ama-1 mRNA was expressed in both mock- and drh-3 RNAi-treated animals (lanes 1 and 2, respectively). RNAi-mediated suppressions of transcripts from other target genes used in this study, rad-51, clk-2, and hus-1, were also confirmed by RT-PCR (Fig. 3B,C).
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It is well known that genetic mutations of RecQ-like WRN DNA helicase, which have a role in maintaining genome integrity, cause accelerated aging in mammals (Bohr 2005), and that mutations of the C. elegans orthologue WRN-1 and the RecQ-like helicase RCQ-5 lead to shortened life span of adult worms (Jeong et al. 2003; Lee et al. 2004). Thus, the influence of drh-3 RNAi on the life span of adult animals was examined. Comparison of the life span of the mock-treated nematode population and that of the drh-3(RNAi) population was compared, but no statistically significant difference was detected, as shown in Fig. 4C (P values are 0.22 for the logrank test and 0.41 for the t-test). However, a significant life span reduction in the control daf-16(RNAi) population was observed, that is, the mean life span was reduced to 52% of the mock-treated population as reported previously (Lin et al. 2001). This also suggests that continuous RNAi treatment worked properly in the present experiments.
Sensitivity of F1 progeny from drh-3 RNAi-treated nematodes to X-ray irradiation and camptothecin
To elucidate the potential involvement of drh-3 in the DNA damage response, the viability of F1 progeny from drh-3(RNAi) P0 animals after X-ray- or camptothecin-induced DNA damage was examined. As previously observed in our screens, the hatching rate of the progeny from drh-3(RNAi) animals was significantly decreased after X-ray irradiation in a dose-dependent manner compared with that of the F1 progeny from control animals (Fig. 5A). The progeny from rad-51(RNAi) animals was highly sensitive to X-ray irradiation, which is consistent with observations by others (Takanami et al. 2000; Rinaldo et al. 2002). The sensitivity of F1 progeny from RNAi-treated P0 animals after exposure to camptothecin was tested. Camptothecin causes DNA double-strand breaks via inhibiting topoisomerase I-catalyzed reactions in cells. As was the case for X-ray irradiation, the progenies from both drh-3(RNAi) and rad-51(RNAi) animals were highly sensitive to camptothecin treatment (Fig. 5B). In addition, no significant differences in sensitivity to UV irradiation or hydroxyurea (HU) treatment between mock- and drh-3 RNAi-treated progenies were found (data not shown).
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Drh-3 RNAi caused embryonic lethality and hypersensitivity to X-ray- or camptothecin-induced DNA damage. Since it has been reported that depletion of nematode RAD-51 causes impaired meiotic recombination, which in turn induces chromosomal aberrations in germ cell nuclei as well as in the above two phenotypes (Takanami et al. 2000; Rinaldo et al. 2002), we investigated germ cell nuclei in gonads of drh-3(RNAi) animals. Typically, six bivalent chromosomes in a diakinesis oocyte nuclei were observed in the proximal region of gonads in mock-treated animals by DNA staining (Fig. 6Aa,c); however, aberrantly aggregated chromosomes were observed in diakinesis oocyte nuclei of drh-3(RNAi) animals (Fig. 6Ab,d), suggesting impaired chromosome segregation. Similar aberrant chromosomal morphologies in oocyte nuclei were detected in gonads of rad-51(RNAi) animals (Fig. 6Ae) as previously reported (Takanami et al. 2000; Rinaldo et al. 2002). Since the oocytes showed abnormal chromosomal morphology in drh-3(RNAi) animals, we examined the early cell cycle division of the fertilized eggs. We observed 4',6'-diamidino-2-phenylindole hydrochloride (DAPI)-stained dividing nuclei in developing early embryos and found abnormally segregated chromosomes (i.e. chromosome bridge) in drh-3(RNAi) embryos (Fig. 6Bb), but not in mock-treated animals (Fig. 6Ba).
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Influence of drh-3 RNAi on checkpoint responses
Many studies have suggested the biological importance of checkpoint responses to aberrant replication fork arrest or DNA damage because dysfunction of the checkpoint response makes the genome unstable and can lead to apoptotic cell death or unregulated cell proliferation. In C. elegans, both checkpoint pathways for aberrant replication arrest and DNA damage are conserved. Typical checkpoint responses can be seen in mitotic germ cells in the distal arm of the gonad where cell proliferation occurs exclusively in the adult nematode (Stergiou & Hengartner 2004). Some checkpoint mutants exhibit increased sensitivity to X-ray irradiation and other DNA-damaging agents, resulting in a reduced reproductive capacity and high embryonic lethality rates as in drh-3(RNAi) animals. Therefore, we examined the checkpoint response to both replication arrests by HU + and DNA damage caused by X-ray irradiation in drh-3(RNAi) animals according to the method of Gartner et al. (2004). The mitotic nuclei in the distal arm of the gonad of each animal were visualized by DNA staining, and the number of nuclei was scored. For the replication checkpoint response, mock-treated animals or drh-3 RNAi- or clk-2 RNAi-treated animals were exposed to 25 mM HU for 24 h after which mitotic germ cell nuclei were observed in their gonads. CLK-2 was found to play a crucial role in the checkpoint response in germ-line proliferation, and clk-2 mutations are known to cause checkpoint defects, for example, continuous cell cycle progression in the distal mitotic zone upon DNA damage (Ahmed et al. 2001). The results shown in Fig. 7A indicate that the checkpoint response occurred normally in mitotic germ cells in both drh-3(RNAi) and mock-treated animals after HU-induced replication arrest. The number of mitotic nuclei was reduced (Fig. 7B), and the size of nuclei was enlarged due to arresting cell proliferation by an activated checkpoint response in both treated animal groups (Fig. 7A). A significant reduction of mitotic germ cells was not detected in gonads from clk-2(RNAi) animals, presumably due to dysfunction of the checkpoint response (Fig. 7B). The DNA replication checkpoint has several functions including retardation of the cell cycle, stabilization of replication forks and inhibition of replication initiation. It should be noted that the latter two reactions in drh-3(RNAi) animals remain to be elucidated, although HU-induced cell cycle arrest occurs normally. The DNA-damage-induced checkpoint response of drh-3(RNAi) animals was also examined together with hus-1(RNAi). Hus-1 encodes a subunit of the 9-1-1 complex that is required for sensing DNA damage and plays a role in the DNA damage checkpoint in C. elegans (Hofmann et al. 2002). A significant decrease in the number of nuclei was observed in gonads from drh-3(RNAi) and mock control animals, but not in gonads from hus-1(RNAi) animals (Fig. 8B). These results clearly indicate that DRH-3 is not involved in the checkpoint responses in C. elegans germ cells.
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Discussion |
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TopAbstractIntroductionResultsDiscussionExperimental proceduresReferences |
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In the present study, we clarified the phenotypic features of C. elegans in which the drh-3 transcripts were suppressed. The increased sensitivity of the drh-3(RNAi) progeny to DNA damaging agents that cause DNA double-strand breaks was confirmed in this study using both X-ray irradiation and camptothecin treatment (Fig. 5), suggesting that drh-3 may play a role in maintaining genome integrity in nematode germ-lines. In addition, we found that drh-3 RNAi-induced embryonic lethality (Fig. 4A) and arrest of embryogenesis in the progeny (Fig. 6C) as well as reduced reproductive capacity at 25 °C (Fig. 4B), induced chromosomal abnormalities in diakinesis oocytes (Fig. 6A), and aberrantly segregated chromosomes (chromosome bridge) in developing early embryos (Fig. 6B) in the DRH-3-depleted animals. These findings clearly indicate that the drh-3 gene is essential for the development of germ-lines in C. elegans, and suggest that embryonic lethality shown in Fig. 4A is probably caused by impaired chromosomal integrity induced by DRH-3 depletion.
DRH-3-depleted animals shared several specific phenotypes with rad-51(RNAi) animals, for example, embryonic lethality (Fig. 4A), X-ray sensitivity (Fig. 4A), and aberrant chromosomes (Fig. 6A) in germ cells. RAD-51 is required both for meiotic homologous recombination and DNA double-strand break repair in germ-lines (Rinaldo et al. 2002). Thus far, we have no evidence that DRH-3 is implicated in the same processes as RAD-51, except for phenotypic resemblance in the RNAi-treated animals.
Drh-3(RNAi) animals exhibited defects similar to those reported in the mutants with defects in DNA repair and checkpoint response. For example, reduction of reproductive capacity and X-ray sensitivity were reported in DNA damage checkpoint-defect mrt-2 and hus-1 mutants (Ahmed & Hodgkin 2000; Hofmann et al. 2002). However, in drh-3(RNAi) worms, the checkpoint responses to aberrant replication arrest by HU and X-ray-induced DNA damages were normal in mitotic germ-lines (Figs 7 and 8). This excludes the possibility that DRH-3 is involved in the checkpoint pathways. In addition, mutations of the RecQ-like DNA helicase WRN-1, which acts in DNA repair, caused shortened life span in C. elegans as well as in humans (Lee et al. 2004; Bohr 2005); however, this was not the case in drh-3(RNAi) animals (Fig. 4C). Taken together, these findings indicate that DRH-3 plays a role in germ-line development by maintaining the chromosomal integrity in C. elegans.
Sequence analysis showed that drh-3 encodes an RNA helicase-like protein that shares significant homology with two C. elegans Dicer-like proteins, DRH-1 and DRH-2 (Fig. 1). Both DRH proteins and Dicer protein (DCR-1) have been shown to play an essential role in several RNAi pathways in C. elegans (Knight & Bass 2001; Tabara et al. 2002), and this raises the possibility that DRH-3 could be implicated in RNAi. Recently, Duchaine et al. (2006) identified the DRH-3 protein as one of the interacting proteins of nematode DCR-1 in a co-purification experiment coupled with mass spectrometry. They showed that drh-3 is required for germ-line RNAi and for the production of several small RNA species with unknown function in C. elegans. They also observed abnormal nuclear morphology in oocytes and impaired chromosomal segregation in somatic nuclei of the drh-3-deleted strain, which is consistent with our observations (Fig. 6A,B). In this study, we observed approximately threefold increase in the levels of drh-3 transcripts during the maturation of larvae to adults (Fig. 2), probably suggesting a potential role of DRH-3 in germ-line development in the gonad of an adult hermaphrodite.
This raises the fundamental question why the RNAi factor DRH-3 is involved in both maintenance of chromosomal integrity and RNAi. While the exact mechanisms involved are still largely unknown, several lines of evidence suggest the existence of RNAi-mediated regulation of chromosomes in several species (Provost et al. 2002; Volpe et al. 2002; Fukagawa et al. 2004; Mochizuki & Gorovsky 2005; Tsukioka et al. 2006). Dicer plays a key role in RNAi processes and appears to be implicated in the silencing of centromeric repeats to ensure accurate chromosome segregation in fission yeast (Provost et al. 2002; Volpe et al. 2002) and vertebrate cells (Fukagawa et al. 2004). This evidence suggests that DRH-3 may be involved in a similar RNAi-mediated process required for faithful chromosome segregation in C. elegans together with DCR-1. However, we cannot exclude a possibility that some small RNA species generated by DRH-3-containing RNAi complex may regulate the expression of proteins required for chromosome segregation. In addition, it has been thought that RNA helicases exclusively play a crucial role in RNAi (Jaronczyk et al. 2005) but DNA helicases do not. However, this notion must be modified because rat RecQ-like DNA helicase RecQ1 has recently been identified as a factor of the Piwi-interacting RNA complex (Lau et al. 2006). Because most of RecQ-like DNA helicases function in the maintenance of genome integrity, there might be an unknown mechanistic link between the DNA repair pathway and RNAi.
Sequence analysis (Fig. 1) revealed that drh-3 encodes the third member of the C. elegans DRHs, and it appears to play a role in RNAi with DCR-1 (Tabara et al. 2002; Duchaine et al. 2006). Several studies suggested functional conservation and diversification of Dicer proteins (Jaronczyk et al. 2005). Caenorhabditis elegans, Schizosaccharomyces pombe and vertebrates encode only one Dicer protein. Drosphila melanogaster have two Dicer proteins (Dcr1 and Dcr2) with unique functions, and the Arabidopsis thaliana genome encodes four Dicer genes, at least three of which have unique functions (Jaronczyk et al. 2005). It remains unclear whether the biological functions of DRHs are conserved among the species. Three Dicer-related proteins (DRH-1, DRH-2 and DRH-3) in C. elegans and three mammalian proteins DDX58, IFIH1 and LGP2 have been identified in eukaryotes, with the three DRH proteins playing a role in RNAi pathways (Tabara et al. 2002; Duchaine et al. 2006) and the three mammalian proteins appearing to play a role in dsRNA-induced signal transduction in innate immunity upon viral infection (Yoneyama et al. 2004, 2005). Further analysis of these proteins is required to elucidate their involvement in RNAi with binding to mammalian Dicer, as in the case of C. elegans DRH proteins. Mph1p is a sole Dicer-like protein in S. cerevisiae (Fig. 1) and belongs to the DRH-related Hef/Mph1p/FANCM subfamily. This subfamily contains human FANCM or archea Hef helicase in addition to Mph1p and many of these members have been shown to play a role in DNA repair processes but not in RNA metabolism. These observations may indicate functional and genetic diversifications of Dicer-related proteins during evolution.
In summary, we have clearly shown that the drh-3 gene encoding a novel Dicer-like protein is implicated in the maintenance of chromosome integrity for germ-line development in C. elegans, but not in checkpoint responses or life span regulation. Because of a possible role for DRH-3 in RNAi, it will be important to biochemically reconstitute a putative RNAi complex containing DRH-3 to clarify interactions with other proteins, including Dicer, so as to further understand the detailed molecular functions of DRH-3 in RNAi and maintenance of chromosomal integrity.
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Experimental procedures |
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Amino acid sequences of proteins were obtained for cluster analysis via the Internet at the WormBase <http://www.wormbase.org/> and NCBI sites <http://www.ncbi.nlm.nih.gov/>. These sequences were aligned by CLUSTALW (version 1.8.3) and a phylogenetic tree was drawn using TREEVIEW software (version 1.6.6). Multiple alignment of representative sequences shown in Fig. 1A was done by CLUSTALX (version 1.8) for the sequence alignment display. To obtain a proper alignment, the N-terminal sequences of nematode DCR-1 and human FANCM (residues 1-700) containing the helicase domain were used.
Caenorhabditis elegans strains and culture procedures
The C. elegans wild-type strain Bristol N2 (kindly provided by Dr I. Katsura, National Institute of Genetics, Japan) was used in this study. Animals were maintained on nematode growth medium (NGM) agar plates seeded with E. coli OP50 strain as described (Harada et al. 2007). Culture of nematodes was carried out at 20 °C unless otherwise specified.
Construction of recombinant DNA
The fragment of cDNA or genomic DNA corresponding to the following gene was cloned into the blunted EcoRI site of the dsRNA expression vector pDP129.36 (gifted from Dr A. Fire, Stanford University School of Medicine, Palo Alto, CA). The cDNA fragments corresponding to drh-3 (D2005.5) and rad-51 (Y43C5A.6) gene were amplified from the phage cDNA clones yk331a2 and yk401c3, respectively, by PCR using the Expand High FidelityPLUS PCR system (Roche Diagnosis K.K., Tokyo, Japan) and the primer set yk5'-F (5'-TGGCGGCCGCTCTAGAACTAGTGGATC-3') and yk3'-SmaR (5'-TTCCCGGGTGAATTGTAATACGACTCACTATAGGGCG-3'). The cDNA fragments of clk-2 (C07H6.6) and daf-16 (R13H8.1) were isolated from the plasmid clone yk1249e03 and yk1044b11, respectively, by PCR with the primer set ME-735FW (5'-GGATGTTGCCTTTACTTCTA-3') and ME-1250RV (5'-TGTGGGAGGTTTTTTCTCTA-3'). These cDNA clones were kindly provided by Dr Y. Kohara (National Institute of Genetics). The 1.1-kb DNA fragment of the entire hus-1 (H26D21.1) gene was amplified from C. elegans genomic DNA (N2 strain) using the primer set hus1-SmaF (5'-GGGGATGAAGTTCAGTTCTCTTCTCCAAGATAC-3') and hus1-SmaR (5'-GGGTTAATCCGAAACATTTCCAACAATATACG-3'). DNA primers were purchased from Sigma Genosys (St. Louis, MO) and Proligo LLC (Boulder, CO). The amplified DNA was blunted with a Blunting kit (TaKaRa, Kyoto, Japan) and phosphorylated by T4 polynucleotide kinase (New England BioLab, Ipswich, MA) for cloning. The nucleotide sequences of the resultant recombinant clones for dsRNA expression were determined using a dye-terminator cycle sequencing kit (GE Healthcare Bio-Sciences, Piscataway, NJ) and an automated DNA sequencer (model 377XL, Applied Biosystems, Foster City, CA). Data were assembled and analyzed using AutoAssembler (version 1.4.0, Applied Biosystems) and Genetyx MAC (version 9, Software Development, Co., Tokyo, Japan), respectively.
Feeding RNAi
RNAi by feeding was performed for drh-3, rad-51, hus-1, clk-2 and daf-16 genes as described with modifications (Timmons et al. 2001; Ohkumo et al. 2006). Briefly, several colonies of the HT115(DE3) E. coli strain (gift from Dr I. Katsura, National Institute of Genetics) containing the recombinant pDP129.36 DNA were mixed and grown overnight in 2xYT or LB medium containing 100 µg/mL ampicillin with shaking at 37 °C. Thirty microliters of the culture was added and spread onto a NGM agar plate for RNAi (i.e. RNAi plate) in a Petri dish 6 cm in diameter, containing 2 mM isopropyl ß-D-thiogalactopyranoside and 100 µg/mL ampicillin, and incubated at 37 °C for 18 h. The next day, P0 animals at the fourth larval (L4) to young adult stages were placed on the plate and fed the recombinant E. coli strain expressing dsRNA specific to the target gene. The transformant containing a pPD129.36 vector alone was used as control bacteria for mock RNAi treatment.
RT-PCR analysis of gene expression
The expressions of drh-3 and rad-51 transcripts following RNAi at 20 °C or 25 °C were examined by RT-PCR using the primer sets D1-RTF (5'-ACAGCAGACTTCACAAGAACAACAATTGAC-3') and D1-RTR (5'-CAAGTACTTACTCACCTTTCGCTTCGATTC-3'), corresponding to nucleotides 2427–2456 and 2956–2985 of the drh-3 cDNA sequence (accession number: NM_059760), and rad51-RTF2 (5'-CAGAAAGCTGAAAAGATAATGAAAGAAGCC-3') and rad51-RTR2 (5'-TTCCTTTTCGAAGGTACAATCTGGTAGTAG-3'), corresponding to nucleotides 366–395 and 1051–1080 of the rad-51 cDNA sequence (NM_001028294), respectively. The expressions of hus-1 and clk-2 transcripts were examined by RT-PCR using the following primer sets hus1-RTf (5'-CGTGTTCTGCCCGGCGGACACTGTTATCTG-3') and hus1-RTr (5'-GATTTGAGATGATATTCATTTTGGCACGTG-3'), corresponding to nucleotides 282–311 and 733–762 of the hus-1 cDNA sequence (NM_058802), and clk2-RTf (5' CATTCACAATGCTGCTGATGGAATGGGTGC-3') and clk2-RTr (5'-TCAAACAGTTCATCCATAAGATCTCCAGGC-3'), corresponding to nucleotides 1919–1948 and 2444–2473 of the clk-2 cDNA sequence (NM_066249), respectively. As a control, ama-1 transcripts were examined using the primers ama-1F2 (5'-ACGCATGTCAGTGGCTCATGTCGAGT-3') and ama-1R2 (5'-CGACCTTCTTTCCATCATTCATCGGATC-3'), corresponding to nucleotides 87–112 and 493–520 of the ama-1 cDNA sequence (NM_068122), respectively. RT-PCR was performed as described below. For RT-PCR analysis of RNAi-treated animals, L1 larvae that were fed dsRNA-expressing bacteria and the resultant adult animals were collected and washed with distilled water. For the analysis of drh-3 mRNA expression during larval development, synchronized animals at each developmental stage (i.e. L1–L2, L3–L4, young adult and adult) were used. Total RNA was extracted with vigorous shaking for 5–10 min at 4 °C using the TRIzol® reagent (Invitrogen, Carlsbad, CA) and glass beads (G-8772, Sigma). Isolated RNA was treated with RQ1 RNase-free DNase (Promega, Madison, WI) and subjected to cDNA synthesis using the ReverTra Ace-
-® kit (Toyobo, Tokyo, Japan) according to the manufacturer's instructions. Each 4-µL aliquot of reaction mixture was added to the tubes containing either an RNAi target gene or ama-1 specific primers. After 30–40 cycles of amplification, the PCR products were separated on 2% agarose gels.
Measurement of hatching rate
The viability of F1 progeny laid by drh-3 RNAi- or rad-51 RNAi-treated P0 worms was monitored by scoring the hatching rate of fertilized eggs as described (Ohkumo et al. 2006). Five L4-stage animals were transferred and grown on an RNAi plate seeded with dsRNA-expressing bacteria in order to lay eggs for 24 h. Subsequently, the animals were transferred onto a fresh RNAi plate containing bacteria for RNAi to lay eggs during the following 24 h. This process was repeated once. At about 12 h after P0 removal, the number of hatched larvae and dead eggs on each plate were scored under a stereomicroscope (MZ FLIII, Leica, Wetzlar, Germany) to determine the hatching rate of fertilized eggs laid in each 24-h culture. Measurement was performed in triplicate and the experiment was repeated at least twice.
Measurement of reproductive capacity
The average number of eggs laid by each drh-3 RNAi- or rad-51 RNAi-treated animal per generation was monitored and the measurements were performed up to the eighth generation at both 20 °C and 25 °C. An L4 animal maintained at 25 °C was placed on a bacterial lawn producing dsRNA to the target gene and allowed to lay eggs for 48 h at the same temperature. When the animals were kept at 20 °C and after laying eggs for 48 h, they were further moved onto a fresh RNAi plate seeded with bacteria for RNAi to lay eggs for 24 h. The total number of eggs laid by a worm during 48 h at 25 °C or during 72 h at 20 °C was determined. Measurement was performed using six independent animals treated by RNAi in each generation, and the mean value of the number of laid eggs per animal was scored.
Determination of life span
The life span assay was performed as described (Harada et al. 2007) with modifications for RNAi. In brief, approximately 40 synchronized L4 worms grown on an RNAi plate with bacteria containing a vector alone were transferred onto four fresh RNAi plates (ten worms per plate) containing a dsRNA-expressing bacterial lawn and 25 µM 5-fluoro-2'-deoxyuridine (FUdR, Sigma). FUdR was used to prevent the contamination of parent worms with offspring. We have confirmed that there was no significant difference in life span of animals cultured with and without FUdR under our experimental conditions (data not shown). After placing the animals on the RNAi plates (life span = 0), the adult animals were checked for viability under the stereomicroscope every day as described (Harada et al. 2007). It is noted that the amount of bacteria for RNAi in the plate was sufficient for maintaining ten adult worms. For statistical analyses, the software package JMP IN5.1.2J (SAS Institute Inc., Cary, NC, USA) was used. The means and standard errors of the life spans for worms in each group were calculated and significant differences in life span in each nematode population were analyzed by the Kaplan–Meier method and Student's t-test (significance set at P < 0.05).
X-ray sensitivity assay
Four young adult worms were fed with bacteria expressing dsRNA to the target gene on an RNAi plate for 12 h and transferred onto a fresh RNAi plate (dish 3 cm in diameter) with a bacterial lawn for RNAi. The animals were irradiated with X-ray (Radioflex 320CG, Rigaku, Tokyo, Japan) at the ratio of 2 Gy/min and transferred onto a fresh RNAi plate with bacteria for RNAi. Animals were cultured for 24 h to lay eggs and then removed. After 36 h, hatching rates of laid eggs were determined. The assay was performed in triplicate. X-ray doses were determined with a condenser R-meter (model 570, Victoreen Instrument, Moedling, Austria).
Camptothecin sensitivity assay
Young adult worms were fed with bacteria producing dsRNA for the drh-3 or rad-51 gene or a control bacteria for 24 h and suspended in 1 mL of M9 buffer (0.6% Na2HPO4, 0.3% KH2PO4, 0.5% NaCl, and 1 mM MgSO4). RNAi-treated animals were collected by centrifugation, suspended in M9 buffer and then transferred into 0.2-mL tubes containing 100 µL of M9 buffer with 0.5% dimethyl sulfoxide (Sigma) in the presence or absence of indicated concentrations of camptothecin (Wako Pure Chemicals, Osaka, Japan). After a 2-h incubation at 20 °C, treated animals were placed onto a fresh RNAi plate for 30 min, three animals were placed onto a fresh RNAi plate to feed RNAi bacteria, allowed to lay eggs for 24 h and then removed to determine the hatching rate of the progeny. The assay was performed in duplicate and repeated twice.
DNA staining of germ-lines and embryos
DNA staining of germ cells and embryos was performed as follows. For analysis of germ cell nuclei in gonads, L4 animals fed with dsRNA-expressing bacteria for 50 h were transferred into a 5-µL drop of M9 buffer on a MAS-coated glass slide (S-9441, Matsunami, Osaka, Japan) for dissection. Slides were kept at –80 °C for 15 min and soaked in a fixative (ethanol : acetic acid = 3 : 1) for 1 h. Air-dried slides were soaked in 50 ng/mL DAPI (Sigma) for 10 min and successively washed in distilled water. Air-dried slides were mounted with 5 µL of 2.5% (w/v) 1,4-diazabicyclo-[2,2,2]octane (DABCO) in 90% glycerol–10% PBS (pH 8.6) on a glass cover slip and sealed with nail polish. DAPI-stained nuclei were examined under an Olympus BX50 DIC microscope using appropriate filters. Fluorescent images were captured with an equipped CCD camera (DP70, Olympus, Tokyo, Japan) and processed by DP manager (Olympus) and Adobe Photoshop (version 7.0, Adobe Systems, Inc., San Jose, CA) software. For analysis of zygotic embryos, P0 animals were fed with dsRNA-expressing bacteria for 24 h and moved onto a fresh RNAi plate with bacterial lawn for RNAi to lay eggs for 24 h. After removing P0 animals, the progeny was cultured for 36 h and the unhatched eggs that remained on the plate were collected by centrifugation with 1 mL of M9 buffer. The unhatched eggs were fixed with 0.4 mL of fixative at 4 °C for 30 min and collected by centrifugation. After washing with M9 buffer, 3 µL of egg suspension was mounted on a glass slide with 2 µL of DABCO solution containing 0.25 mg/mL DAPI and a glass cover slip. The slides were sealed with nail polish to observe nuclei under the microscope, and fluorescent and DIC images for more than 20 eggs per sample were captured with a CCD camera for further analysis.
Analysis of checkpoint response in mitotic germ-lines
Experiments were performed as described (Gartner et al. 2004) with the following modifications. For analysis of replication checkpoint, L4-to-young adult-stage animals were fed with bacteria producing dsRNA on an RNAi plate. After 24 h, RNAi-treated worms were placed onto a fresh RNAi plate with bacteria for RNAi in the presence or absence of 25 mM HU (Sigma) for 24 h. HU solution was added onto a bacterial lawn in the plate, and the plate was dried and used for the experiment within a single day. Treated animals were dissected for DAPI-stained gonad samples as described above. Fluorescent images of gonad at three different levels were captured to score the number of mitotic germ cell nuclei. More than eight gonads per group were examined and the experiment was repeated twice. Checkpoint response to DNA damage caused by X-ray irradiation was examined as follows. In brief, hatched larvae were fed with bacteria expressing dsRNA and grown young adults were irradiated with indicated doses of X-ray as described above. Animals were cultured on a fresh RNAi plate with dsRNA-expressing bacteria for 9 h followed by dissection. Mitotic nuclei in gonads were scored as described above. More than four gonads per group were examined. The experiment was repeated using mock-treated, drh-3(RNAi) and rad-51(RNAi) animals.
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Acknowledgements |
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* Correspondence: E-mail: eki{at}eco.tut.ac.jp
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Received: 3 September 2006
Accepted: 29 May 2007
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