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Genes to Cells (2008) 13, 609-621. doi:10.1111/j.1365-2443.2008.01192.x
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Possible involvement of caspase-7 in cell cycle progression at mitosis

 Toshiaki Hashimoto1, Lisa Yamauchi1, Tony Hunter2, Ushio Kikkawa1 and Shinji Kamada1,*

1 Biosignal Research Center, Kobe University, 1-1 Rokkodai-cho, Nada-ku, Kobe 657-8501, Japan
2 Molecular and Cell Biology Laboratory, The Salk Institute, 10010 North Torrey Pines Road, La Jolla, CA 92037, USA


   Abstract
Top
 Abstract
Introduction
 Results
 Discussion
 Experimental procedures
 References
 
Caspases are suggested to play essential roles not only in apoptotic but also in non-apoptotic functions. However, the contribution of caspases to the cell cycle regulation is unclear. Here we found that caspases including caspase-3, caspase-7, caspase-8 and caspase-9 were activated during mitosis. Chemically synthesized caspase inhibitors delayed mitotic progression and induced accumulation of mitotic cells, which exhibited abnormal chromatin condensation and incomplete chromosome segregation. Furthermore, knockdown of caspase-7 by using small interfering RNAs resulted in the inhibition of cell proliferation, but knockdown of other caspases did not show a significant effect on cell growth. The expression of short hairpin RNA directed against caspase-7 induced the cell cycle arrest at mitosis, which was rescued by the re-expression of caspase-7 containing silent mutations at the target site for the short hairpin RNA. These results revealed that caspase-7 has a novel role during cell cycle progression at mitosis.


   Introduction
Top
 Abstract
 Introduction
Results
 Discussion
 Experimental procedures
 References
 
Cell cycle progression is monitored by signaling pathways at key transitions to ensure the successful completion of upstream events prior to proceeding to the next phase. These regulatory pathways are referred to as cell cycle checkpoints functioning at G1/S, S, G2/M and M phases (Musacchio & Hardwick 2002; Kastan & Bartek 2004). Checkpoint signaling induces not only cell cycle arrest to repair cellular damage but also activation of apoptotic machineries if cellular damages were not properly repaired. Defects in cell cycle checkpoints often result in gene mutations, chromosome deterioration and aneuploidy, all of which contribute to tumorigenesis (Jallepalli & Lengauer 2001; Musacchio & Hardwick 2002; Bharadwaj & Yu 2004; Rajagopalan & Lengauer 2004).

Caspases, a family of cysteine proteases, are required for apoptosis execution and cytokine maturation (Cryns & Yuan 1998; Thornberry & Lazebnik 1998; Earnshaw et al. 1999). Caspases are zymogens that consist of three regions, an N-terminal prodomain, a large subunit and a small subunit. At least 11 human caspase subtypes have been identified that are divided into three groups. The caspases with a large prodomain are referred to as inflammatory caspases including caspase-1, caspase-4 and caspase-5, and as initiator caspases of apoptosis including caspase-2, caspase-8, caspase-9 and caspase-10. The caspases with a short prodomain are referred to as effector caspases of apoptosis, and include caspase-3, caspase-6 and caspase-7. The activation of effector caspases is catalyzed through proteolytic processing by initiator caspases, whereas the activation of large prodomain containing caspases is suggested to occur in a protein complex generated by the binding through the prodomains. The large prodomains have characteristic structures, such as the death effector domain (DED) and the caspase recruitment domain (CARD). Caspase-8 and caspase-10 contain two tandem DEDs and are activated in the death-inducing signaling complex (DISC). The CARD is found in caspase-1, caspase-2, caspase-4, caspase-5 and caspase-9, and plays an essential role for the interaction with other CARD-containing proteins leading to the activation of these caspases.

Although it is suggested that caspases are required for non-apoptotic functions such as cell proliferation and differentiation (Newton & Strasser 2003; Woo et al. 2003; Huh et al. 2004; Fernando et al. 2005; Kuranaga et al. 2006; Siegel 2006; Lamkanfi et al. 2007), and a role of caspase-3 during mitosis is pointed out (Swe & Sit 2000; Yan et al. 2001; Hsu et al. 2006), the physiological importance of caspases in mitosis and the contribution to cell proliferation have been unclear. We report here that several caspases including caspase-7 are activated from the late G1 to M phase in an apoptosis-independent manner, and small interfering RNAs (siRNAs) and short hairpin RNA (shRNA) directed towards caspase-7 prevented cell proliferation through the cell cycle arrest at mitotic phase. These results showed a novel function of caspase-7 in the regulation of mitosis.


   Results
Top
 Abstract
 Introduction
 Results
Discussion
 Experimental procedures
 References
 
Mitotic cells were stained with anti-active caspase-3 antibodies

The antibodies specific for active caspase-3 have been used for detection of caspase-3 activation in situ (Yan et al. 2001; Fernando et al. 2005; Kamada et al. 2005a,b). When apoptotic HepG2 cells were stained with an anti-active caspase-3 antibody, normally proliferating HepG2 cells were used as a negative control for staining. Unexpectedly, we found that a small but distinct population of normal HepG2 cells (approximately 5%) was stained with the anti-active caspase-3 antibody, which recognizes the newly exposed C-terminus of the caspase-3-p17 subunit generated during proteolytic activation of procaspase-3 (Fig. 1A). The nuclei of the cells stained with the anti-active caspase-3 antibody were condensed, but not fragmented, suggesting that these cells were in mitotic phase of cell cycle.


Figure 1
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  		Figure 1  Mitotic HepG2 cells are stained with anti-active caspase-3 antibodies. (A) After fixation and permeabilization, normally proliferating HepG2 cells were incubated with the anti-active caspase-3 pAb (G7481) that recognizes the newly exposed C-terminus of caspase-3-p17, and the TXRD-labeled secondary antibody, followed by staining the nuclei with Hoechst 33342. (B) After fixation and permeabilization, normally proliferating HepG2 cells were incubated with the anti-active caspase-3 pAb (G7481, left panel) or the anti-active caspase-3 polyclonal antibody (2622, right panel) that recognizes the newly exposed N-terminus of caspase-3-p12, and the anti-{alpha}-tubulin monoclonal antibody, and TXRD- and FITC-labeled secondary antibodies, followed by staining the nuclei with Hoechst 33342. The exposure time was at least 5 times longer than that for apoptotic HepG2 cells.

 
To confirm whether the cells stained with the anti-active caspase-3 antibody are mitotic cells, normal HepG2 cells were co-stained with anti-active caspase-3 antibodies recognizing p17 or p12, and an anti-{alpha}-tubulin monoclonal antibody (Fig. 1B). {alpha}- and β-tubulins are the major building blocks of microtubules, which form the mitotic spindle and function as structural and motile elements during mitosis. As shown in Fig. 1B, HepG2 cells were not stained during interphase with either anti-active caspase-3 antibody specific for p17 or p12, whereas cells at all stages of mitosis were stained with these antibodies, implying that caspase-3 is activated specifically at mitosis.

Caspase activation during cell cycle progression

To validate the immunofluorescence staining data, we next set out to detect the caspase-3-p17 subunit in cells in mitosis by immunoblot analysis. After synchronization of HeLa cells at late G1 with a double-thymidine block following by release into the cell cycle (Fig. 2A), cells were collected at each time point and analyzed by immunoblotting. The active form of caspase-3 as well as caspase-9 was detected at 10, 12 and 14 h after release from the thymidine block (Fig. 2B). Furthermore, active forms of caspase-7 and caspase-8 were detected from late G1 to M phase and the levels of active caspase-7 and caspase-8 were elevated at M phase. The cleavage of caspase-3 and caspase-7 substrates, such as poly-ADP-ribose polymerase (PARP), lamin B1 and PKC{delta}, was observed at M phase, supporting that activated caspases have proteolytic activities during M phase. Although the activation of caspases was detected at M phase, apoptotic cells with sub-2n DNA content were not significantly increased at M phase (Fig. 2A). The activation of caspases was transient, because active caspases were not detected in normally proliferating cells, and no apparent decrease in the level of procaspases was observed in M phase cells despite the appearance of active caspases, indicating that the activation of caspases is much lower than in the apoptotic cells (Fig. 2B). It seems that caspase activation in normal cells is tightly regulated, and that full activation of caspases that would lead to apoptotic cell death, is avoided. These results suggest that caspase-3, caspase-7, caspase-8 and caspase-9 are potentially involved in the regulation at M phase.


Figure 2
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  		Figure 2  Caspases and caspase substrates are cleaved during cell cycle progression. (A) Cell cycle profiles of synchronized HeLa cells. Cells were arrested at late G1 phase with a double-thymidine block, followed by release to enter S phase were analyzed by flow cytometry. (B) Immunobot analysis of synchronized HeLa cells. Cell lysates from HeLa cells at each cell cycle point as described in (A) were subjected to SDS-PAGE, followed by immunoblotting with antibodies as indicated. A, The lysate from apoptotic HeLa cells as a positive control. N, The lysate from normally proliferating HeLa cells as a negative control. (C) Delay of cell cycle progression at mitotic phase by the caspase inhibitor. HeLa cells were arrested at late G1 phase with a double-thymidine block, followed by release to enter the cell cycle. At 7 h after the release, 0.5% DMSO or 200 µM Z-Asp-CH2-DCB was added to the culture medium, and the DNA content of cells at each point was analyzed by flow cytometry.

 
To analyze the role of caspases in the regulation of mitosis, we monitored cell cycle progression in cells treated with caspase inhibitor. The synthetic peptide-based inhibitor for caspases were dissolved in dimethyl sulfoxide (DMSO) and added to the culture medium. Since DMSO has some toxicity for cells, we determined the highest concentration of DMSO that could be added to culture medium without affecting proliferation. DMSO at 3% had no effect on cell proliferation of HepG2 cells and the compound at 0.6% was inert for HeLa cells (data not shown). Therefore, DMSO at these concentrations was used for each cell line as a vehicle for the reagents in the following experiments. After synchronization in late G1 with a double-thymidine block, HeLa cells were released to enter the cell cycle and a broad-spectrum caspase inhibitor (Z-Asp-CH2-DCB) was added to culture medium at 7 h after release from the late G1 arrest (Fig. 2C). The entry into the next G1 of the caspase inhibitor-treated cells was delayed about 2 h compared with the DMSO-treated cells, suggesting that caspase activation plays a positive role in mitotic progression.

Inhibition of caspase activities induced mitotic arrest

Next, we examined the effect of caspase inhibitors on nuclear morphology to clarify the mechanisms of mitotic progression (Fig. 3). We monitored the nuclear morphology of HeLa cells, in which histone H2B-GFP fusion protein is stably expressed (HeLa-H2B-GFP) to visualize chromatin (Kanda et al. 1998), using confocal microscopy after treatment with Z-Asp-CH2-DCB. Although the control cells treated with DMSO transited through mitosis within 120 min (Fig. 3A,B, and Supplementary Movies S1–S3), the nuclei of caspase inhibitor-treated cells showed mitotic abnormalities, including failure of chromosome alignment on the metaphase plate (Fig. 3C,D) or arrest at metaphase (Fig. 3E, and Supplementary Movies S4 and S5). Similar abnormalities were observed when HeLa-H2B-GFP cells were treated with other caspase inhibitors, including Boc-Asp-(OMe)-FMK, Boc-Asp(Obzl)-CMK (data not shown) and caspase-3/caspase-7 inhibitor in which the hydrophobic region of the signal peptide of Kaposi fibroblast growth factor (K-FGF) was N-terminally fused to the peptides to confer cell-permeability, Ac-AAVALLPAVLLALLAP-DEVD-CHO (Supplementary Movie S6). Since many cells with condensed, but not fragmented, nuclei showing mitotic rather than apoptotic were observed after treatment with caspase inhibitors (Supplementary Movies S4–S6), the number of cells containing condensed nuclei was counted after treatment of HepG2 cells with Z-Asp-CH2-DCB. As shown in Fig. 3F, the ratio of caspase inhibitor-treated cells with condensed nuclei was increased as compared with control cells. These results clearly show that the inhibition of caspase activities perturbs mitotic progression and suggest that caspases play an important role(s) in mitosis.


Figure 3
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  		Figure 3  Caspase inhibitor treatment induces abnormal chromatin morphology during mitosis. (A–E) After treatment of HeLa-H2B-GFP cells with 0.6% DMSO (A and B) or 300 µM Z-Asp-CH2-DCB (C–E) for 2 days, nuclear morphology was observed with confocal microscopy for 60 min (A and B) or 120 min (C–E) at 1 min interval. Typical nuclear morphology of each cell at 0, 30 and 60 min (A and B) or at 0, 60 and 120 min (C–E) from supplementary Movies S2–S5 was represented. (F) HepG2 cells were cultured in the presence of 200 µM Z-Asp-CH2-DCB for different days, followed by staining the nuclei with Hoechst 33342. The percentage of the cells showing condensed nuclei to the total cells was measured. The data (mean ± SD) were obtained from at least three independent experiments.

 
Knockdown of caspase-7 prevented cell proliferation and induced mitotic arrest

To study further the importance of caspases in the regulation of mitosis and to clarify which caspase is essential for cell proliferation, we used siRNA (Elbashir et al. 2001) to deplete caspases in HepG2 cells (Fig. 4). At 2 days after transfection of siRNAs for each caspase, the levels of the target caspases were decreased significantly, whereas the level of {alpha}-tubulin was unaffected (Fig. 4A). The same number of cells transfected with siRNAs for caspases were re-seeded at 4 h after transfection, and cell viability was monitored by WST-1 assay. Although HepG2 cells transfected with single siRNAs for caspase-3, caspase-4, caspase-8 and caspase-9 proliferated as well as the mock- or control siRNA-transfected cells, caspase-7 siRNA (C7) effectively prevented cell proliferation (Fig. 4B). Furthermore, when combined with caspase-7, but not caspase-3, siRNA, siRNAs for caspase-3, caspase-4, caspase-8 and caspase-9 prevented cell proliferation (Fig. 4C,D). To exclude off-target effects of siRNA for caspase-7 (C7), three other siRNAs for caspase-7 (C7–226, C7–478 and C7–940) were designed and used to deplete caspase-7. As shown in Fig. 4E,F, these siRNAs also decreased the expression level of caspase-7 and inhibited cell proliferation. These results strongly suggest that caspase-7 plays an essential role in cell proliferation.


Figure 4
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  		Figure 4  Knockdown of caspase-7 prevents cell proliferation. (A) and (B) After transfection of siRNAs for control, caspase-3 (C3), caspase-4 (C4), caspase-7 (C7), caspase-8 (C8) and caspase-9 (C9) into HepG2 cells, the expression levels of each caspase were determined by immunoblot analysis at day 2 (A), and cell viability was determined by WST-1 assay at days 1, 2 and 3 (B). The data were obtained from three independent experiments. (C) siRNAs for control, caspase-3, and caspase-3 in combination with siRNAs for caspase-4, caspase-7, caspase-8 and caspase-9 were transfected into HepG2 cells, and cell proliferation was analyzed as described in (B). (D) siRNAs for control, caspase-7, and caspase-7 in combination with siRNAs for caspase-3, caspase-4, caspase-8 and caspase-9 were transfected into HepG2 cells, and cell proliferation was analyzed as described in (B). (E) and (F) After transfection of siRNAs for control and caspase-7 (C7-226, C7-478 and C7-940) into HepG2 cells, the expression level of caspase-7 was determined by immunoblot analysis at day 2 (E), and cell proliferation was analyzed (F) as described in (B). The data were obtained from nine independent experiments. The data (mean ± SD) in (B), (C), (D) and (F) indicate as the relative values to the data at day 1 after transfection in each siRNA transfectant.

 
To further confirm the effects of caspase-7 knockdown in cell cycle progression, pKD-Caspase7-v1, which expresses caspase-7 shRNA, was transfected into HeLa-H2B-GFP cells and the nuclear morphology was monitored for 2 days (Fig. 5). Transiently transfected pKD-Caspase7-v1 decreased the level of exogenously expressed caspase-7-GFP (Fig. 5A) and endogenous caspase-7 (to < 35%) (Fig. 5B). To ascertain the effects of shRNA for caspase-7 on cell proliferation, a fivefold excess of shRNA expression plasmid over DsRed expression plasmid was transfected into HeLa-H2B-GFP cells, and the DsRed-positive cells were monitored as shRNAs expressing cells under time-lapse fluorescence microscopy. When pKD-NegCon-v1, which expressed control shRNA, were transfected, most cells proliferated normally (79.3% of DsRed-positive cells) and a minor population of transfected cells died during interphase (17.2% of DsRed-positive cells) (Fig. 5C,D, and Supplementary Movie S7). However, most of the pKD-Caspase7-v1 transfected cells were arrested and died during mitotic phase (57.1% of DsRed-positive cells) (Fig. 5C,D, and Supplementary Movie S8), consistent with a contribution of caspase-7 to the cell cycle progression at mitosis.


Figure 5
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  		Figure 5  Knockdown of caspase-7 by shRNA induces cell cycle arrest. (A) Inhibition of transiently expressed caspase-7-GFP by shRNA for caspase-7. 293T cells (3 x 105 cells) were transfected with 1 µg of pcasp7-Wt-GFP in combination with pKD-NegCon-v1 or pKD-Caspase7-v1 as indicated amount of DNA using Lipofectamine. At 2 days after transfection, cells were harvested and cell lysates were fractionated with SDS-PAGE, followed by immunoblotting with antibodies as indicated. (B) Decrease of endogenous caspase-7 by shRNA for caspase-7. 293T cells (1.3 x 106 cells) were transfected with 1.7 µg of pKD-NegCon-v1 or pKD-Caspase7-v1 using Nucleofector Kit V. At 2 days after transfection, cells were harvested and cell lysates were fractionated with SDS-PAGE, followed by immunoblotting with antibodies as indicated. (C) and (D) Nuclear morphology of shRNA transfected cells. HeLa-H2B-GFP cells were seeded at a density of 1 x 105 cells per 35-mm glass bottom dish. After 12 h, cells were transfected with 0.5 µg of pDsRed-Monomer-C1 (BD Biosciences) with 2.5 µg of pKD-NegCon-v1 or pKD-Caspase7-v1. At 48 h after transfection, DsRed-positive cells were observed with a time-lapse fluorescence microscope for 48 h at 15 min intervals. DsRed-positive cells were divided into proliferated or dead cells judging from the nuclear morphology during time-lapse observation. Dead cells having fragmented nuclei were identified during M phase which shows chromosome alignment at metaphase plate or interphase which shows normal nuclei, and the number of each type of cells and the percentages to the total DsRed-positive cells were shown in (C). Typical features of cells transfected with shRNA expression plasmids for control or caspase-7 were shown in (D) (see Supplementary Movies S7 and S8).

 
Next, we carried out rescue experiments in which mutant caspase-7 having silent mutations in the target site for caspase-7 shRNA was co-expressed with pKD-Caspase7-v1 (Fig. 6). After determination of the target site sequence for pKD-Caspase7-v1 by sequencing, we constructed two types of mutant caspase-7; one is wild type caspase-7 with silent mutations (pHis-casp7W-sh1) and the other one is inactive caspase-7 with silent mutations in which active site cysteine was replaced with serine (pHis-casp7M-sh1) (Fig. 6A). Transiently transfected pKD-Caspase7-v1 decreased the level of exogenously expressed caspase-7-GFP, whereas the level of exogenously expressed His-tagged caspase-7 with silent mutations was not affected by pKD-Caspase7-v1 (Fig. 6B). To assess the effects of pHis-casp7W-sh1 or pHis-casp7M-sh1 in caspase-7 knockdown cells, pKD-Caspase7-v1 and DsRed expression plasmid were transfected into HeLa-H2B-GFP cells with pHis-casp7W-sh1 or pHis-casp7M-sh1, and the DsRed-positive cells were monitored as shRNA and His-tagged caspase-7 expressing cells under time-lapse fluorescent microscopy (Fig. 6C). Transfection of pHis-casp7W-sh1 caused an increase in the percentage of proliferating cells as compared with pcDNA3.1His-transfected cells (30.7% increase) accompanied by a decrease in the percentage of dead cells during M phase (23.7% decrease). However, these effects were significantly reduced when inactive caspase-7 with silent mutations (pHis-casp7M-sh1) was re-expressed. These results confirmed that the inhibition of cell proliferation by caspase-7 shRNA is dependent on the depletion of caspase-7, and strongly suggest that caspase-7 plays an essential role in cell cycle progression at mitotic phase.


Figure 6
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  		Figure 6  Rescue of caspase-7 knockdown by transient expression of caspase-7 with silent mutations. (A) Diagram showing the constructions of pHis-casp7W-sh1 and pHis-casp7M-sh1. Silent mutations are shown in bold. Asterisk shows the active center (QACRG) of caspase-7. (B) No decrease in the level of transiently expressed His-tagged caspase-7 carrying silent mutations by shRNA for caspase-7. 293T cells (1.2 x 105 cells) were transfected with 0.25 µg of pcasp7-Wt-GFP, pHis-casp7-W-sh1 or pHis-casp7-M-sh1 in combination with 0.75 µg of pKD-NegCon-v1 or pKD-Caspase7-v1 using FuGENE 6. At 2 days after transfection, cells were harvested and cell lysates were fractionated with SDS-PAGE, followed by immunoblotting with antibodies as indicated. (C) Effects of transient expression of caspase-7 with silent mutations in caspase-7 shRNA transfected cells. HeLa-H2B-GFP cells were seeded at a density of 5 x 104 cells per 35-mm glass bottom dish. After 12 h, cells were transfected with 0.1 µg of pDsRed-Monomer-C1 and 0.45 µg of pKD-Caspase7-v1 with 0.45 µg of pcDNA3.1/His A, pHis-casp7W-sh1 or pHis-casp7M-sh1. At 24 h after transfection, DsRed-positive cells were observed with a time-lapse fluorescence microscope for 48 h at 15 min intervals. DsRed-positive cells were classified into proliferating or dead cells judging from the nuclear morphology during time-lapse observation. Dead cells having fragmented nuclei were identified during M phase which shows chromosome alignment at metaphase plate or interphase which shows normal nuclei, and the number of each type of cells and the percentages of the total DsRed-positive cells analyzed are shown.

 

   Discussion
Top
 Abstract
 Introduction
 Results
 Discussion
Experimental procedures
 References
 
Among caspases, caspase-7 has not been focused on its physiological function, because its substrate specificity is not distinct from caspase-3 (Cryns & Yuan 1998; Thornberry & Lazebnik 1998; Earnshaw et al. 1999) and no obvious abnormality is found in caspase-7–/– mice (Lakhani et al. 2006). Here we show for the first time the physiological role of caspase-7 in the regulation of cell cycle progression at mitosis. It has been suggested that caspase-3 is involved in the cell cycle regulation at mitotic phase; active caspase-3 fragment is immunostained in the proliferative region in rat brain (Yan et al. 2001), caspase-3 is up-regulated just prior to mitosis (Hsu et al. 2006), and peptide-based caspase-3 inhibitors at high concentrations induce cell death at late mitosis (Swe & Sit 2000). However, the contribution of caspase-3 to the regulation of mitosis is not clear, and the peptide-based caspase-3 inhibitor prevents both caspase-3 and caspase-7 activities. We observed the activation of caspase-3 as well as that of caspase-7, caspase-8 and caspase-9 at mitotic phase by immunostaining and immunoblotting. The possibility of the off-target effects of the broad-spectrum caspase inhibitor (Z-Asp-CH2-DCB) employed on the inhibitions of other cysteine proteases could not be excluded (Schotte et al. 1999; Misaghi et al. 2006), as relatively high concentrations of the caspase inhibitor was needed for inhibition of cell cycle progression. We consider, however, that caspase-7 plays an essential role in cell proliferation for the following reasons; (i) other peptide-based caspase inhibitors, Boc-Asp-(OMe)-FMK and Boc-Asp(Obzl)-CMK, had the same effects on cell cycle progression, (ii) lower concentrations of a caspase-3 and caspase-7 specific inhibitor induced mitotic arrest, (iii) knockdown of caspase-7 using siRNAs and shRNA resulted in the inhibition of cell proliferation and the arrest at mitotic phase, (iv) the arrest at mitotic phase with caspase-7 shRNA was rescued by re-expression of caspase-7 with silent mutations. Like multiple caspases are activated through amplification loop in apoptotic cells, caspase-3 might be activated contingently after caspase-7 activation at mitotic phase. Alternatively, caspase-3 would also contribute to the cleavage of substrates to regulate cell cycle progression at mitotic phase in collaboration with, but less effectively than caspase-7.

Since caspase-7 is expressed as a latent proenzyme, it is important to elucidate how caspase-7 is activated during the cell cycle and whether the apoptotic caspase activation machinery is also involved in this activation. Caspase-8 and caspase-9 function as initiator caspases to activate caspase-3 and caspase-7 in apoptotic cells (Cryns & Yuan 1998; Thornberry & Lazebnik 1998; Earnshaw et al. 1999). However, knockdown of caspase-8 or caspase-9 by siRNAs transfection did not show apparent inhibition of cell proliferation. Further studies are required to elucidate the potential contribution of caspase-8 and/or caspase-9 to the cell cycle regulation through caspase-7 activation, since caspase-8 or caspase-9 still remained in the knockdown cells, even small amounts, in our experiments. Interestingly, it was reported that active caspase-8 was translocated into the nuclei followed by activated caspase-7, which in turn cleaved the chromosomal passenger proteins, CENP-C and INCENP, in apoptotic cells (Faragher et al. 2007). Although we did not confirm the nuclear localization of active caspase-7, a portion of caspase-7 may be localized in nuclei, which contributes to the regulation of cell cycle progression at mitosis.

Caspase-3–/– mice are born at lower frequencies, and are smaller than their littermates (Kuida et al. 1996), whereas caspase-7–/– mice are born with no abnormality (Lakhani et al. 2006). Since double knockout mice lacking caspase-3 and caspase-7 die immediately after birth (Lakhani et al. 2006), the postnatal development of caspase-3–/– or caspase-7–/– mice may be rescued by caspase-7 or caspase-3, respectively. It has been reported that a deficiency in caspase-3 or caspase-9 can induce activation of caspase-6 and caspase-7, or caspase-2 and caspase-6, respectively, in knockout mice (Zheng et al. 2000). In addition, the depletion of caspase-3 by siRNA induces a compensatory elevation in caspase-7 level (Wurzer et al. 2003). Furthermore, we observed an elevation of caspase-6 in HeLa and HepG2 cells in which caspase-3 and caspase-7 were depleted by the expression of shRNAs for caspase-3 and caspase-7 using a lentiviral expression system (L. Yamauchi & S. Kamada, unpublished data). Therefore, cells deficient in caspase-7 would not necessarily be expected to show an impairment of cell proliferation because of compensatory functions of other caspases that are not observed when caspase-7 are acutely depleted from cells. A series of our data demonstrated that caspase-7 contributed to the cell cycle regulation at mitotic phase, but these results do not necessarily exclude the possible contribution of other caspases, including caspase-3, caspase-8 and caspase-9. It will be necessary to address these issues in the next step.

Activation of caspases by proteolytic cleavage is an irreversible reaction. Therefore, the mechanisms for silencing caspase activities are critical for prevention of apoptotic cell death during cell cycle progression. Inhibitor of apoptosis proteins (IAPs) such as cIAP1, cIAP2 and XIAP function not only as caspase inhibitors but also as E3 ubiquitin ligases (Salvesen & Duckett 2002; Riedl & Shi 2004). Recently, several groups reported that activated caspase-3 and caspase-7 can be ubiquitinated by IAPs, such as cIAP2 and XIAP, and degraded by the proteasome (Huang et al. 2000; Suzuki et al. 2001). Furthermore, it was reported that over-expression of cIAP1 suppressed cell proliferation, and that although cIAP1 was localized in nuclei during interphase, cIAP1 was released into the cytoplasm early in mitosis, then re-localized in nuclei in late anaphase and in telophase (Samuel et al. 2005). Therefore, IAPs may function as regulators for caspases during cell cycle progression by their activities as caspase inhibitors and/or E3 ubiquitin ligases.

It has been reported that caspases, including caspase-3/caspase-7, caspase-8 and caspase-9, are constitutively activated in some human tumor cells (Yang et al. 2003), and that IAPs are important for maintaining tumor cell survival (Sasaki et al. 2000; Fulda et al. 2002; Schimmer et al. 2004). We observed the activation of caspase-3, caspase-7, caspase-8 and caspase-9 in the cells in an apoptosis-independent manner. On the other hand, many proteins cleaved by caspases during apoptosis have been identified that function in cell cycle progression. For example, the Cdk inhibitors p21 and p27, cyclin E and Rb regulate the transition of cell cycle from G1 to S phases, and Bub1, BubR1, Scc1/Rad 21, CENP-C and INCENP are involved in M phase progression (Fischer et al. 2003; Kim et al. 2005; Faragher et al. 2007; Lüthi & Martin 2007). These proteins are essential for cell cycle checkpoints and dysfunction of these proteins has been suggested to contribute to tumorigenesis (Musacchio & Hardwick 2002; Bharadwaj & Yu 2004; Kastan & Bartek 2004). Therefore, caspases activated in tumor cells may contribute to loss of cell cycle checkpoints and facilitate the rapid proliferation of these tumor cells. It will be interesting to elucidate the contribution of apoptosis-independent caspase activation and cleavage of their substrates to tumorigenesis. The identification of the critical caspase-7 targets needed for mitotic progression may provide clues to address these issues.


   Experimental procedures
Top
 Abstract
 Introduction
 Results
 Discussion
 Experimental procedures
References
 
Antibodies

Preparation of anti-active caspase-3 polyclonal antibody (2622), which recognizes the N-terminus of caspase-3-p12, was described previously (Kamada et al. 2005a). Anti-active caspase-3 pAb (G7481), which recognizes the C-terminus of caspase-3-p17, was obtained from Promega (Madison, WI); anti-caspase-3 polyclonal antibody (9662), anti-cleaved caspase-7 polyclonal antibody (9491), anti-caspase-8 polyclonal antibody (9764), and anti-PARP polyclonal antibody (9542) were from Cell Signaling (Danvers, MA); anti-caspase-4 monoclonal antibody (M029-3) and anti-caspase-9 monoclonal antibody (M054-3) were from Medical and Biological Laboratories (Nagoya, Japan); anti-caspase-7/MCH-3 monoclonal antibody (610812) and anti-DsRed polyclonal antibody (8376-1) were from BD Biosciences (Rockville, MD); anti-lamin B1 polyclonal antibody (sc-6217) and anti-PKC {delta} polyclonal antibody (sc-937) were from Santa Cruz Biotechnology (Santa Cruz, CA); anti-{alpha}-tubulin monoclonal antibody (T-5168) was from Sigma (St Louis, MO).

Cell culture, synchronization and apoptosis induction

HepG2 (a hepatocellular carcinoma line) cells were cultured in RPMI 1640 medium with 10% fetal bovine serum (FBS). HeLa (a cervical carcinoma line, clone D98AH2) and HeLa-H2B-green fluorescent protein (GFP) (Kanda et al. 1998) cells were cultured in DMEM supplemented with 10% FBS. For late G1 phase synchronization, HeLa cells were seeded at a density of 4 x 105 cells per 60-mm dish and cultured for 24 h. After exposure to 2.5 mM thymidine for 18 h, cells were washed with phosphate-buffered saline (PBS) 3 times and incubated in fresh medium for 10 h, and then exposed to 2.5 mM thymidine again for 14 h. To release the cells from the late G1 arrest, medium containing thymidine was removed, and the cells were washed with PBS 3 times and incubated in fresh medium for different times. For induction of apoptosis, HeLa cells were treated with 1 µg/mL agonistic anti-Fas antibody (CH-11; Medical and Biological Laboratories) for 4 h. Transfection was performed using Lipofectamine (Life Technologies, Gaithersburg, MD) or Nucleofector Kit V (Amaxa Biosystems, Koeln, Germany) for 293T cells, FuGENE 6 (Roche, Basel, Switzerland) for HeLa-H2B-GFP and 293T cells according to the manufacturer's instructions.

Fluorescence microscopy

For immunofluorescence analysis, cells were fixed with 3.7% formaldehyde in PBS for 10 min, washed with PBS twice, permeabilized in 0.5% Triton X-100 in PBS for 10 min, and washed with PBS twice. Cells were then incubated with primary antibodies in PBS containing 1% bovine serum albumin overnight at 4 °C. After washing with PBS twice, cells were incubated with Texas Red (TXRD)- or fluorescein isothiocyanate (FITC)-labeled secondary antibodies for 10 min at room temperature, and washed with PBS twice. After staining nuclei with 10 µM Hoechst 33342 (Calbiochem, La Jolla, CA), cells were examined under a fluorescence microscope (Leitz Laborlux).

Time-lapse fluorescence microscopy

For time-lapse fluorescence microscopy, HeLa-H2B-GFP cells were plated on a 35-mm glass bottom dish. The medium was replaced with MEM supplemented with 5% fetal calf serum without phenol red, and dishes were placed in a humidified chamber at 37 °C that was mounted on a confocal laser scanning microscope (model FV500; Olympus) or a fluorescence microscope (model BZ-8000; Keyence) with a constant supply of mixed air containing 5% CO2. For analyses with FV500, samples were excited at 488 nm with an argon laser and detected with a 60x oil immersion objective. Image data were obtained automatically every 1 min by using FLUOVIEW software (Olympus). For analyses with BZ-8000, cells were observed with a 60x oil immersion objective lens and image data were obtained automatically every 15 min by using BZ-H1TL software (Keyence). In the original program by the manufacturer, cells are exposed to the intense mercury lamp light intermittently and the cells were driven to apoptosis within the initial 6 h by the physical damage of the light. Therefore, we used the neutral density filters to reduce the light levels to 1.6% that allowed the cells to proliferate during 48 h's observation.

Flow cytometry

HeLa cells (1 x 106 cells) were incubated with 0.5 mL of PI/RNase buffer (BD Bioscience) after fixation with 70% ethanol for 1 h at –20 °C, and DNA content was measured using FACSCaliber (Becton-Dickinson). About 2 x 104 events were analyzed for each sample, and data were plotted using MODIFIT software, and cell cycle profiles were determined by CELLQuest (Becton-Dickinson).

Immunoblot analysis

Cells were lysed in lysis buffer [20 mM Tris–HCl (pH 7.5), 150 mM NaCl, 1% Nonidet P-40, 50 µg/mL phenylmethanesulfonyl fluoride, 5 mM EDTA]. Protein samples were separated by SDS-polyacrylamide gel electrophoresis and blotted onto Immobilon polyvinylidene difluoride membrane (Millipore, Bedford, MA). Each protein was detected using primary antibodies as indicated, horseradish peroxidase-conjugated secondary antibodies, and ECL-plus detection reagent (GE Healthcare, Little Chalfont, UK).

Cell proliferation assay

Cell proliferation reagent WST-1 (Roche) was used as index of cell viability according to the manufacturer's instructions. In brief, HepG2 and HeLa cells were plated at 4 x 103 cells per well in a 96-well plate with 100 µL medium and incubated for different days, followed by addition of WST-1 reagent to medium and additional incubation for 2 h. The cleavage of the tetrazolium salt WST-1 by mitochondrial dehydrogenases was measured by absorbance at 450 nm and at 690 nm as the reference wavelength.

RNA interference

Synthetic double-stranded siRNA for caspase-3, whose target sequence was AAGATCATACATGGAAGCGAA, corresponding to coding nucleotides 54–75 relative to the first nucleotide of the start codon, was designed by Qiagen (Valencia, CA). siRNAs for caspase-4 (Hs_CASP4_5, SI00299558), caspase-8 (Hs_CASP8_7, SI00299593), and caspase-9 (Hs_CASP9_5, SI00299600) were obtained from Qiagen; caspase-7 (sc-29929) and control siRNA (sc-37007) were obtained from Santa Cruz Biotechnology. Synthetic double-stranded siRNAs for caspase-7 were obtained from Japan Bio Services (Saitama, Japan); AAGAACTTTGA TAAAGTGACA (C7-226) corresponding to coding nucleotides 226–246 relative to the first nucleotide of the start codon, AAGGATTTGACAGCCCACTTT (C7-478) corresponding to 478–498, AAGCAATGGGTCACTCATTAA (C7-940) corresponding to 940–960. Synthetic double-stranded siRNAs for caspases were transfected with TransMessenger Transfection Reagent (Qiagen) according to the manufacturer's instructions. In brief, one day before transfection, HepG2 cells were seeded at a density of 1 x 105 cells per well of 24-well plate. In a tube, 1 µg of siRNA duplex was mixed with 100 µL of Buffer EC-R containing 1.6 µL Enhancer R. When two siRNAs were introduced in combination, 0.5 µg of each siRNA was employed. After incubation for 5 min at room temperature, 4 µL of TransMessenger Transfection Reagent was added to the mixture, and then incubated for 10 min at room temperature. After addition of 100 µL DMEM without serum to the tube, the entire mixture was added to the cells and incubated for 4 h. For cell proliferation assay, cells were re-seeded at a density of 5 x 103 cells per well of 96-well plate and cultured for different days. Mammalian caspase-7 shRNA expression plasmid pKD-Caspase7-v1 (62-004) and mammalian negative control shRNA expression plasmid pKD-NegCon-v1 (62-002) were obtained from Upstate (Lake Placid, NY).

Plasmid constructions

Construction of pcasp7-Wt-GFP was described elsewhere (Kamada et al. 2005a). To construct mutant caspase-7 containing six silent mutations between amino acids 213 and 218, a PCR method was used with the primers, 5'-ATACAAGATTCCT GTTGAGGCCGATTTCCTCTTC-3' and 5'-GAAGAG GAAATCGGCCTCAACAGGAATCTTGTAT-3', and wild type caspase-7 cDNA or mutant caspase-7 cDNA containing active site mutation as templates. The resultant fragments were cloned into the BamHI site of pcDNA3.1/His A (Invitrogen) to generate pHis-casp7W-sh1 and pHis-casp7M-sh1.


   Acknowledgements
 
We thank Dr Teru Kanda for HeLa-H2B-GFP cells. This work was supported in part by the 21st Century COE Program of the Ministry of Education, Culture, Sports, Science and Technology of Japan, and by Public Health Service Grants CA82863 and CA14195 from the National Cancer Institute (to T. H.). T. H. is a Frank and Else Schilling American Cancer Society Professor.


   Footnotes
 
Communicated by: Tadashi Yamamoto

* Correspondence: Email: skamada{at}kobe-u.ac.jp


   References
Top
 Abstract
 Introduction
 Results
 Discussion
 Experimental procedures
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Received: 8 January 2008
Accepted: 5 March 2008





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