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FJ
EXPRESS SUMMARY ARTICLE The Full-length version of this article is also available, published online January 5, 2006 as doi:10.1096/fj.05-5020fje. |
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Department of Biochemistry and Molecular Genetics, University of Colorado Health Sciences Center, Aurora, Colorado, USA
1Correspondence: Department of Biochemistry and Molecular Genetics, University of Colorado Health Sciences Center at Fitzsimons, Mail Stop 8101, PO Box 6511, Aurora, CO 80045, USA. E-mail: jessica.tyler{at}uchsc.edu
SPECIFIC AIM
The purpose of our study was to address the function of the histone chaperone Anti-silencing function 1 (ASF1) in metazoan cells by studying its localization and the consequences of RNA interference-mediated knockdown of Drosophila melanogaster ASF1 (dASF1).
PRINCIPAL FINDINGS
1. Loss of dASF1 causes slow cell proliferation
To investigate the function of ASF1, RNA interference (RNAi) was used to reduce levels of dASF1 protein (knockdown) in Drosophila S2 cells in culture. The exposure of cells to double-stranded RNA (dsRNA) homologous to Drosophila ASF1 resulted in the knockdown of ASF1 protein to undetectable levels from day 2 to 8 after dsRNA exposure as measured by Western blot. Knockdown of dASF1 protein levels correlated with almost complete loss of nuclear dASF1 signal by immunofluorescence in all cells on day 2 after dsRNA exposure.
Because yeast deleted for ASF1 grow slowly and Drosophila ASF1 biochemically assembles chromatin during DNA replication, we examined cell proliferation after dASF1 depletion by RNAi. Exposure of cells to ASF1 dsRNA led to a small but reproducible reduction in cell numbers compared with controls beginning 6 days after dsRNA treatment. This decrease in cell number was a result of slow cell proliferation, not increased cell death, because cells lacking dASF1 did not activate apoptosis as measured by an Annexin V binding assay. Furthermore, there were not increased numbers of dead cells in cultures exposed to dASF1 dsRNA as determined by Trypan blue staining. We conclude that the RNAi-mediated knockdown of Drosophila ASF1 in cells in culture results in slow cell proliferation.
2. Loss of dASF1 results in defective DNA replication
We next asked whether the slowing of cell proliferation after dASF1 knockdown corresponds to defective DNA replication. On day 4 after exposure of cells to dASF1 dsRNA there was an increase in the proportion of cells with G1 phase DNA content and with DNA content intermediate between G1 and G2/M as compared with controls, consistent with an accumulation of dASF1 knockdown cells in S phase of the cell cycle. We found that treatment of cells with hydroxyurea, which depletes nucleotide pools, leading to stalled replication, resulted in accumulation of cells with a G1 phase DNA content, consistent with the accumulation of cells with a G1 phase DNA content after dASF1 depletion also being due to a DNA replication defect. Furthermore, we found that passage through S phase was 
2 h slower in cells after dASF1 depletion, as measured directly by BrdU pulse chase analysis. GFP dsRNA exposed cells had a slight cell cycle abnormality with an accumulation of cells in S phase, which may be a result of the activation of RNAi. However, this did not correspond to a slowing of passage through S phase as measured by BrdU pulse chase analysis. These results indicate that dASF1 promotes passage through S-phase and is in agreement with recent data showing that the RNAi-mediated knockdown of the human homologues of ASF1 leads to an accumulation of cells in S phase of the cell cycle.
We then examined DNA replication after knockdown of dASF1 more thoroughly by determining whether replication forks are established when dASF1 is depleted. Cells were analyzed 4 days after RNA exposure because at later time points control cells had begun to reach quiescence, complicating the analysis. Components of the replication fork, including PCNA and the MCM complex, are known to form foci detectable by immunofluorescence during active replication in Drosophila cells. We found that 49% of dASF1 knockdown cells had co-localizing MCM2-7 and PCNA foci, as compared with 18.7% of untreated cells and 21.3% of GFP dsRNA exposed cells. Replication foci vary in appearance depending on whether a replicating cell is in early, middle, or late S phase. The loss of dASF1 did not result in a change in the distribution of cells with replication foci corresponding to early, middle, and late S phase. These data indicate that the loss of dASF1 results in the accumulation of cells at all stages of S phase.
We next examined replication directly in asynchronously growing control and knockdown cells by measuring bromodeoxyuridine (BrdU) incorporation. Treatment of cells with hydroxyurea was used as a positive control to verify that our assay measures a defect in replication. We found that the number of cells incorporating BrdU after dASF1 depletion was not increased over controls to the extent predicted by the number of knockdown cells with replication foci. By graphing the ratio of BrdU-positive cells to the proportion of cells with replication foci, we found that approximately half of the dASF1 knockdown cells with replication foci were not incorporating BrdU. The discrepancy between the proportions of control cells that have replication foci and are BrdU positive is likely to represent the technical differences between these measurements, as cells in S phase at any time during the hour of BrdU exposure incorporate BrdU, while only those cells in S phase at the time of harvest have detectable replication foci. Furthermore, depletion of dASF1 led to a "collapse" of the arc of S phase cells, indicating that a subpopulation of cells in S phase for the entire BrdU exposure period incorporated less BrdU compared with control cells. We conclude that the loss of dASF1 leads to a defect in DNA replication and a slowing of cells in S phase of the cell cycle.
3. dASF1 co-localizes with active replication forks
Because yeast with ASF1 deleted have deregulated transcription, it is possible that the defective replication seen in cells lacking dASF1 is an indirect effect of transcriptional deregulation. Therefore, we asked whether dASF1 plays a direct or indirect role during DNA replication. After removal of soluble dASF1 with Triton-X, we found that chromatin-bound dASF1 forms foci that co-localize with MCM foci in S phase cells (Fig. 1
). We did not detect ASF1 foci in nonreplicating cells. Furthermore, dASF1 co-localized with MCM foci in early, middle, and late stages of S phase. This result is not due to antibody cross-reaction or detection of fluorescence signal outside of the correct channel because co-localizing foci were not seen when slides were exposed to each primary antibody alone and exposure of cells to dASF1 dsRNA abrogated the dASF1 signal. This result suggests that the replication defect occurring in the absence of dASF1 is not due to indirect effects of transcription deregulation but that dASF1 may play a direct role during DNA replication.
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Next, we investigated whether dASF1 localization to replication forks is dependent on active replication. The approach was to disrupt ongoing replication by treatment with hydroxyurea. We found that treatment of asynchronously growing cells with hydroxyurea disrupted dASF1 foci (Fig. 1)
, while MCM foci remained, presumably due to MCM localized to stalled replication forks as previously reported in yeast. We conclude that chromatin-bound dASF1 co-localizes with active, but not stalled, replication forks.
CONCLUSIONS AND SIGNIFICANCE
We find that depletion of Drosophila ASF1 leads to a defect in DNA replication (Fig. 2
). This replication defect is most consistent with a problem with replication elongation, as PCNA is known to be present during replication elongation and cells lacking dASF1 accumulate with PCNA foci. Furthermore, we observe twice as many cells with replication foci when dASF1 is absent as compared with the control cells, yet only slightly more cells lacking dASF1 were BrdU positive as compared with control cells. These data indicate that even though more cells have the replication machinery loaded onto the DNA when dASF1 is absent, there are not more cells undergoing active replication. Finally, the anomalous BrdU arc in cells lacking dASF1 is consistent with a defect in replication elongation, not initiation. Consistent with a role for dASF1 during replication elongation is that yeast with ASF1 deleted has genomic instability during S phase, which may be a consequence of the stalling of replication forks resulting in DNA double-strand breaks and genomic instability.
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Why would the absence of dASF1 lead to defects in replication elongation? Having shown that dASF1 localizes to replication foci, one possibility is that dASF1 assembles the newly-replicated DNA into chromatin in vivo as it does in vitro (Fig. 2)
. There is precedent for chromatin assembly factors influencing DNA replication, as RNAi-mediated knockdown of CAF-1 results in S phase arrest and defects in BrdU incorporation. Alternatively, ASF1 has recently been shown to be a global chromatin disassembly factor in yeast, such that dASF1 may disassemble chromatin prior to DNA replication (Fig. 2)
.
We find that dASF1 co-localizes with active replication forks throughout S phase (Fig. 2)
. This suggests that ASF1 has a general function during DNA replication that is not specific, for example, to replicating heterochromatin late in S phase. Furthermore, this result suggests that the function of ASF1 is coupled to ongoing DNA replication, as would be expected for a protein involved in chromatin assembly and/or disassembly. It will be important to determine the replication fork components that mediate the localization of ASF1 to the replication foci. One obvious candidate is CAF-1, which localizes to replication foci via its interaction with PCNA, and is known to directly bind to ASF1. Another possibility is that replication factor C (RFC) may recruit Asf1 to replication forks in vivo; it has been recently shown that RFC binds to Asf1 in vitro and can recruit Asf1 to DNA in vitro. It is also possible that Asf1 simply localizes to regions of active chromatin assembly in a nonspecific fashion.
Treatment of cells with hydroxyurea disrupts dASF1 foci, suggesting that dASF1 is lost from stalled replication forks (Fig. 2)
. How the loss of chromatin-bound dASF1 from stalled replication forks is regulated remains to be determined. It is possible that ASF1 localization is regulated by the S phase checkpoint, as human and Drosophila ASF1 are phosphorylated by tousled-like kinases (tlks) during S phase of the cell cycle, and this phosphorylation is inhibited by activation of the S phase checkpoint. However, both phosphorylated and nonphosphorylated ASF1 functionally interact with CAF-1, suggesting that checkpoint activation would not cause ASF1 to be lost from replication forks if its localization was via its interaction with CAF-1/PCNA. Recent data show that during treatment with hydroxyurea human ASF1 joins a multichaperone complex in the cytosol and that this process is not checkpoint mediated. It is possible that the same unknown regulatory mechanism that governs ASF1’s localization to the cytosol during replication stress also controls localization of ASF1 to the replication fork.
In summary, we find that Drosophila ASF1 localizes to active replication forks, and that the loss of dASF1 results in defective replication and the accumulation of cells in S phase of the cell cycle. These results suggest that dASF1 may have a direct and important role in the modulation of chromatin structure during DNA replication in vivo.
FOOTNOTES
To read the full text of this article, go to http://www.fasebj.org/cgi/doi/10.1096/fj.05-5020fje;
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