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Infection and Immunity, April 2006, p. 2080-2092, Vol. 74, No. 4
0019-9567/06/$08.00+0     doi:10.1128/IAI.74.4.2080-2092.2006
Copyright © 2006, American Society for Microbiology. All Rights Reserved.

Exposure of Lymphocytes to High Doses of Actinobacillus actinomycetemcomitans Cytolethal Distending Toxin Induces Rapid Onset of Apoptosis-Mediated DNA Fragmentation

Bruce J. Shenker,^1* Donald R. Demuth,² and Ali Zekavat¹

Departments of Pathology,¹ Biochemistry, University of Pennsylvania School of Dental Medicine, Philadelphia, Pennsylvania²

Received 24 October 2005/ Returned for modification 16 November 2005/ Accepted 11 January 2006

Top Abstract Introduction Materials and Methods Results Discussion References

 
We have shown previously that treatment of human lymphocytes with the Actinobacillus actinomycetemcomitans cytolethal distending toxin (Cdt) results in dose-dependent G₂ arrest, followed 24 h later by apoptotic cell death. Here we demonstrated that for Jurkat cells exposed to high concentrations of Cdt (>0.2 ng/ml) there was a dose-dependent increase in the level of S-phase cells and a concomitant decrease in the level of G₂ cells. Fluorescence-activated cell sorter analysis demonstrated that the S-phase cells did not incorporate BrdU and likely represented cells that arrested in G₂ and underwent significant DNA fragmentation. Analysis of the kinetics of the appearance of both S-phase cells and apoptotic cells supported this interpretation. Cells exposed to low doses of toxin exhibited G₂ arrest at 24 h, but at 48 and 72 h there were also decreases in the level of G₂ cells and concomitant increases in the levels of S, G₀/G₁, and sub-G₀ cells; these changes were paralleled by increased numbers of apoptotic cells. Cells exposed to high doses of toxin exhibited these changes 24 to 48 h earlier. We also examined the relationship between G₂ arrest, DNA fragmentation, and activation of the apoptotic cascade. We employed two inhibitors of apoptosis, overexpression of Bcl-2 and the caspase-3 inhibitor zvad. Both inhibitors blocked Cdt-induced apoptosis, Cdt-induced DNA fragmentation, and phosphorylation of the histone H2AX. However, the cells retained the ability to undergo G₂ arrest in the presence of the toxin. Thus, it appears that high doses of Cdt induce rapid onset of DNA degradation resulting from activation of the apoptotic cascade.

Top Abstract Introduction Materials and Methods Results Discussion References

 
It is becoming increasingly evident that perturbation of the cell cycle of host cells is a common target utilized by a number of pathogens. For instance, the vpr genes expressed by lentiviruses, such as human immunodeficiency virus, encode proteins that cause G₂ arrest (20, 28). Similarly, Helicobacter pylori has been shown to inhibit the G₁-to-S transition of gastric epithelial cells (48). Another toxin produced by Haemophilus ducreyi induces fibroblasts to arrest in the G₁ phase of the cell cycle (4). The benefit to the pathogen of inducing cell cycle arrest in host cells is not known, although there are a number of theoretical advantages. For instance, cell cycle arrest might provide improved conditions for pathogen replication. Another possibility is that cell cycle arrest often leads to altered cell function; in this regard, immunologic responses are dependent upon the ability of lymphocytes to proliferate. Therefore, microbial toxins that interfere with this process would induce a form of local and/or systemic immunoparalysis and provide clear advantages for pathogen survival. Thus, it is noteworthy that we and other workers have demonstrated that a number of pathogenic bacteria produce immunoinhibitory agents that appear to act via their ability to subvert lymphocyte cell cycle progression. For instance, Fusobacterium nucleatum and Treponema denticola produce immunoinhibitory proteins that induce G₁-phase arrest of human lymphocytes; in the case of F. nucleatum, the G₁ arrest is associated with the failure to express cyclin D3 and PCNA (26, 45). Recently, the cytolethal distending toxin (Cdt) has been shown to induce G₂ arrest in human T cells and other cell lines (3, 6, 38, 40, 42, 46, 50).

The Cdts are a family of heat-labile protein cytotoxins produced by Actinobacillus actinomcyetemcomitans, Escherichia coli, H. ducreyi, Campylobacter jejuni, Salmonella enterica serovar Typhi, and Shigella species (3, 16, 30, 36, 37, 39-41, 46). Cdt is encoded by three cotranscribed genes designated cdtA, cdtB, and cdtC that encode polypeptides with apparent molecular masses of approximately 24 to 35 kDa (39-41, 46). Moreover, we have recently demonstrated that the Cdt holotoxin consists of Cdt subunits CdtA, CdtB, and CdtC. The Cdt holotoxin appears to function as an AB₂ toxin, in which CdtB is the active (A) unit and the complex consisting of CdtA and CdtC comprises the binding (B) unit (24, 33, 44). Furthermore, it is generally agreed that the active subunit, CdtB, enters the cell, while CdtA and CdtC remain associated with the cell surface (5, 31, 33). Cdts were first characterized on the basis of their ability to cause progressive cellular distension and finally death in some cell lines. We have demonstrated that Cdt is a potent immunotoxin that induces lymphocytes to undergo irreversible G₂ arrest; toxin-induced G₂ cells have been shown to contain elevated levels of the inactive, hyperphosphorylated form of cdk1, suggesting that the these cells are unable to progress into the M phase of the cell cycle due to an inactive cdk1-cyclin B complex (46, 47).

In previous studies we demonstrated not only that treatment of lymphocytes with Cdt results in cell cycle arrest but also that the toxin-treated cells eventually become apoptotic. More recently, not only did we observe that induction of G₂ arrest by Cdt holotoxin was dose dependent, but we also noted that exposure of lymphocytes to high levels of toxin resulted in a reduction in the accumulation of G₂ cells. Therefore, we investigated the relationship between cell cycle arrest, G₂ cell accumulation, apoptosis, and toxin-induced DNA fragmentation. Here we report that high doses of Cdt do indeed result in reduced G₂ cell accumulation; the decrease corresponds both to increases in cells in the sub-G₀, G₀/G₁, and S-phase regions and to an increase in DNA fragmentation. Furthermore, we found that the rapid onset of DNA fragmentation induced by high concentrations of toxin are not the result of direct effects of the toxin but rather are irreversible effects of cell cycle arrest that lead to activation of the apoptotic cascade.

Top Abstract Introduction Materials and Methods Results Discussion References

 
Cell lines. The T-cell leukemia cell line Jurkat (E6-1; American Type Tissue Culture Collection, Manassas, VA) was maintained as previously described (43). Jurkat cells transfected with the bcl-2 gene (Jurkat^bcl-2) and control Jurkat cells transfected with the control pcDNA3 vector (Jurkat^neo) were kindly provided by J. C. Reed (The Burnham Institute, La Jolla, CA). Cells were maintained in RPMI 1640 supplemented with 10% fetal calf serum, 2 mM glutamine, 10 mM HEPES, 100 U/ml penicillin, and 100 µg/ml streptomycin. Cells were harvested in the mid-log growth phase and plated at a concentration of 5 x 10⁵ cells/ml in 24-well tissue culture plates. The cells were exposed to medium or Cdt holotoxin and incubated for 18 h or as indicated below.

Expression and purification of recombinant Cdt holotoxin. Construction and characterization of plasmid pUCAacdtABC^his, which expresses the Cdt holotoxin containing a His tag on the CdtC peptide, have been described previously (43). The plasmid was constructed so that the cdt genes were under control of the lac promoter; all ligation mixtures were transformed into E. coli DH5. Cultures of transformed E. coli were grown in 1liter LB broth and induced with 0.1 mM isopropyl-ß-D-thiogalactopyranoside (IPTG) for 2 h; bacterial cells were harvested, washed, and resuspended in 50 mM Tris (pH 8.0). The cells were frozen overnight, thawed, and sonicated. The His-tagged holotoxin was isolated by nickel affinity chromatography as previously described (43). Briefly, the sonicated bacterial extracts were applied to a histidine binding column (HiTrap Chelating HP; Amersham Biosciences, Uppsala, Sweden). The column was washed and His-tagged proteins eluted with 500 mM imidazole; previous studies have demonstrated that the isolated holotoxin, designated CdtABC, consists of a heterotrimeric complex of CdtA, CdtB, and CdtC (43).

Cell cycle and S-phase analysis. Jurkat cell cultures were set up as described above and incubated for 18 h; 30 min prior to harvest, the cultures received bromodeoxyuridine (BrdU) (10 µg/ml; Sigma Chemical Co., St. Louis, MO). The cells were harvested, washed, fixed in 80% ethanol for 30 min at –20°C, and incubated for 30 min at room temperature in 2 N HCl containing 0.5% (vol/vol) Triton X-100. Cells were then washed in 0.1 M Na₂B₄O₇ (pH 8.5), resuspended in 0.5% (vol/vol) Tween 20 containing 1.0% bovine serum albumin, and stained for 30 min with anti-BrdU monoclonal antibody conjugated to fluorescein isothiocyanate (FITC) (Becton Dickinson Immunocytometry Systems, San Jose, CA). Following washing in Tween 20, the cells were stained with propidium iodide (PI) (2 µg/ml; Sigma Chemical Co.) and analyzed by flow cytometry. FITC and PI fluorescence were excited by an argon laser at 488 nm (250 mW), and fluorescence was measured through 530/30 and 630/22 band-pass filters, respectively. Stained cells were analyzed simultaneously for immunofluorescence (FITC) and cell cycle distribution (PI) using a Becton-Dickinson Facstar^PLUS flow cytometer; electronic compensation was used to remove spectral overlap. Immunofluorescence data were collected using log amplification, and PI fluorescence data were obtained using linear amplification; a minimum of 15,000 events were collected for each sample.

Analysis of DNA fragmentation and cell cycle. DNA fragmentation in Cdt-treated Jurkat cells was measured using the terminal deoxynucleotidyltransferase-mediated dUTP-biotin nick end labeling (TUNEL) assay (in situ cell death detection kit; Boehringer Mannheim, Indianapolis, IN) in conjunction with Hoechst 33342 to measure cell cycle progression. Jurkat cell cultures were prepared as described above; at the end of the incubation period cells were centrifuged, resuspended in 1 ml of freshly prepared 4% formaldehyde, and vortexed gently. After incubation for 30 min at room temperature, the cells were washed with phosphate-buffered saline (PBS) and permeabilized in 0.1% Triton X-100 for 2 min at 4°C. The cells were then washed with PBS and incubated in a solution containing FITC-labeled nucleotide and terminal deoxynucleotidyltransferase according to the manufacturer's specifications. Following the final wash, the cells were resuspended in PBS containing Hoechst 33342 (5 µg/ml; Molecular Probes) and analyzed by flow cytometry. FITC fluorescence was excited by an argon laser at 488 nm (250 mW), and emission was measured through a 530/30-nm band-pass filter. Hoeschst 3342 fluorescence was excited by a second laser operating in UV mode (330 nm; 50 mW), and emission was measured through a 424/44-nm band-pass filter.

Analysis of caspase-3 activation. Caspase-3 activation was monitored by flow cytometry. Jurkat cell cultures were prepared as described above; at the end of the 18-h incubation cells were treated with the fluorescent inhibitor FAM-DEVD-FMK according to the manufacturer's specifications (CaspTag fluorescein caspase activity kit; Chemicon, Temecula, CA). Cells were analyzed by flow cytometry as previously described (47).

Measurement of mitochondrial permeability transition (MPT). Jurkat cells were treated with medium or Cdt as described above for 18 h. Development of the permeability phase transition was monitored by simultaneously assessing the transmembrane potential (_m) and generation of superoxide anions using 40 nM 3,3'-dihexyloxacarbocyanine [DiOC₆(3)] (Molecular Probes, Eugene, OR) and 2 µM hydroethidine (Molecular Probes), respectively (1, 19). Fluorescence was measured after the cells were stained with the fluorochromes for 15 min at 37°C. The probes were excited with a laser at 488 nm (250 mW), and emission was monitored through a 530/30-nm band-pass filter for DiOC₆(3) and a 575/26-nm band-pass filter for ethidium; the latter compound is the fluorescent product of hydroethidine oxidation by superoxide anion (19). Logarithmic amplification was used to detect fluorescence; at least 10,000 cells were analyzed per sample.

Analysis of H2AX phosphorylation. Jurkat^neo and Jurkat^bcl-2 cells were treated with medium or Cdt holotoxin (200 pg/ml) as described above. After 4 h, the cells were harvested and fixed in paraformaldehyde. The cells were then permeabilized and stained with anti-phospho H2AX as recommended by the manufacturer (Upstate USA, Inc., Lake Placid, NY). Samples were then analyzed by fluorescence-activated cell sorting (FACS) as described above; at least 10,000 cells were analyzed per sample.

Top Abstract Introduction Materials and Methods Results Discussion References

 
We demonstrated previously that the Cdt holotoxin is composed of three truncated subunits, CdtA, CdtB, and CdtC, and that both normal human lymphocytes activated by mitogen and the T-cell leukemia cell line Jurkat were exquisitely sensitive to the effects of this toxin (43). As shown in Fig. 1, for Jurkat cells exposed to various amounts of Cdt holotoxin there was a dose-dependent increase in the level of cells in the G₂ phase of the cell cycle. For cells exposed to 0.32 pg/ml Cdt 18.3% of the cells were G₂ cells, while 46.6% G₂ cells were observed in cultures treated with 200 pg/ml toxin; this compares to 9% G₂ cells in the control (medium alone) population. Surprisingly, for cells exposed to CdtABC concentrations of >0.2 ng/ml there was a dose-dependent decrease in the percentage of G₂ cells; for instance, we observed 28.3% and 6.0% G₂ cells in cultures treated with 1 and 25 ng/ml toxin, respectively. Figure 2A shows the cell cycle distribution for cells exposed to medium, to a low dose of CdtABC (0.05 ng/ml), which induced G₂ accumulation, and to a high concentration of CdtABC (5 ng/ml), which resulted in a significant reduction in the level of G₂-phase cells. Exposure to the low dose of toxin resulted in a fourfold increase in the level of G₂ cells (43.9%) compared to the level observed in control cultures. In contrast, the high CdtABC dose resulted in a 90% reduction in the level of G₂ cells (4.7%) along with a concomitant increase in the level of S-phase cells (91.6%) compared to cells exposed to the low dose of toxin.

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FIG. 1. Effect of Cdt holotoxin on lymphocyte G2 arrest and DNA fragmentation. Jurkat cells were treated with various concentrations of Cdt holotoxin for 18 h. The cells were then analyzed by flow cytometry to determine both the cell cycle distribution and the presence of DNA fragmentation as described in Methods and Materials. The percentage of G2 cells (•) and the percentage of cells exhibiting DNA fragmentation () are plotted versus Cdt concentration; the data are means ± standard deviations for three experiments. For control cells exposed to medium 9.1% of the cells were in the G2 phase; 4.9% of the control cells exhibited DNA fragmentation.

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FIG. 2. Comparison of the effects of Cdt on cell cycle arrest, BrdU incorporation, and DNA fragmentation. Jurkat cells were treated with medium (control) or a low dose (0.05 ng/ml) or high dose (5 ng/ml) of Cdt for 18 h. The cells were then analyzed by flow cytometry to determine the cell cycle distribution using propidium iodide (A) and BrdU incorporation in order to directly assess the percentage of cells in the S phase (B) and DNA fragmentation (TUNEL) along with the cell cycle (Hoechst fluorescence) (C). The percentages of cells in the phases of the cell cycle (based on propidium iodide fluorescence) are shown in panel A. The percentages of S-phase cells determined by BrdU incorporation are indicated in panel B; cells that exhibited fluorescence greater than the fluorescence observed in controls (indicated by a line) were considered to be positive. The percentages of cells exhibiting DNA fragmentation are indicated in panel C; fluorescence values greater than the fluorescence observed in cells treated with the control antibody (indicated by a line) were considered to be positive. The results are representative of at least three experiments.

The apparent accumulation of S-phase cells in the presence of high doses of Cdt holotoxin could have resulted from two different mechanisms. For instance, DNA degradation leading to a loss of total DNA in G₂ cells could have resulted in cells shifting into the S-phase region. Alternatively, activation of a different cell cycle regulatory pathway leading to S-phase arrest could have accounted for these observations. To examine these possibilities, CdtABC-treated Jurkat cells were assessed for DNA fragmentation using the TUNEL assay, and S-phase cells were quantitated, independent of the total DNA content, by monitoring BrdU incorporation. We also monitored the accumulation of G₂- and S-phase cells over time in the presence of both low and high CdtABC doses. As shown in Fig. 1, there was a dose-dependent increase in the percentage of cells exhibiting DNA fragmentation. DNA fragmentation was minimal at toxin levels (0.00032 to 0.2 ng/ml) that induced increased accumulation of G₂ cells. However, the percentage of cells exhibiting DNA fragmentation increased at toxin concentrations of >0.2 ng/ml; these doses also resulted in decreases in the levels of G₂ cells and concomitant increases in the levels of S-phase cells. In the presence of a low toxin dose, 8.9% of the cells exhibited DNA fragmentation, while we observed fragmented DNA in 69.7% of the cells exposed to high concentrations of CdtABC (Fig. 2C); for comparison, only 2.5% of the control Jurkat cells exhibited DNA fragmentation. It should be noted that the highest density of cells exhibiting DNA fragmentation was in the sub-G₂ region (Fig. 2C), which most likely corresponded to the S-phase population identified in Fig. 2A. In contrast to the results of assessment of DNA fragmentation, analysis of BrdU incorporation indicated that the percentages of S-phase cells in cultures treated with a low dose of toxin and in control cells (36.9% and 30.7%, respectively) corresponded to the percentages of S-phase cells determined using propidium iodide (Fig. 2A). However, cells exposed to the high dose of Cdt holotoxin failed to incorporate BrdU, and only 4.2% of these cell were in the S phase (Fig. 2B); this percentage of S-phase cells was significantly lower than the percentage determined using propidium iodide (Fig. 2A).

Our results suggest that the decreased accumulation of G₂ cells at high concentrations of CdtABC was due to DNA degradation. To further demonstrate this, we exposed cells to medium alone and low and high CdtABC doses and monitored the cells both for cell cycle progression using propidium iodide and for DNA fragmentation using the TUNEL assay at 24, 48, and 72 h. For Jurkat cells treated with CdtABC for 24 h 52.4% (low dose) and 15.8% (high dose) of the cells were G₂ cells (Fig. 3); 26.3% and 57.8% of the cells exposed to the low and high toxin doses, respectively, were identified as S-phase cells. The cells in the sub-G₀ region accounted for 2.4% (control), 5.7% (low Cdt dose), and 14.4% (high Cdt dose) of the cells. At 48 h, for cells exposed to the low dose of toxin there was an increase in the level of S-phase cells (39.2%) and there was a reduction in the level of G₂ cells (30.5%); for cells exposed to the high dose of toxin there was a decline in the level of S-phase cells (26.6%) and there was a significant increase in the percentage of cells in the sub-G₀ region (51.4%). At 72 h, 39.7% and 38.2% of the cells treated with the low dose of CdtABC were in the sub-G₀ and S phases, respectively; in comparison, cells treated with the high Cdt dose were almost entirely in the sub-G₀ phase (89.2%). It should be noted that the cell cycle distribution of the control cells was essentially unchanged throughout this period.

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FIG. 3. Kinetics of Cdt-induced cell cycle arrest. Jurkat cells were treated with Cdt or with medium for 24, 48, or 72 h. Following the incubation period, cells were stained with propidium iodide and analyzed by flow cytometry to determine cell cycle distribution. The percentages of cells in the different phases of the cell cycle are indicated. The results are representative of at least three experiments.

The kinetics of toxin-induced DNA fragmentation are shown in Fig. 4. The percentage of control cells exhibiting DNA fragmentation at 24, 48, or 72 h was minimal (<4%). Treatment with 0.05 ng/ml CdtABC resulted in a time-dependent increase in the level of cells exhibiting DNA fragmentation, as follows: 8.2% at 24 h, 61% at 48 h, and 72.1% at 72 h. In contrast, after exposure to 5.0 ng/ml CdtABC 51% of the cells exhibited DNA fragmentation within 24 h; the percentage of such cells increased to 77% at 48 h and to 81% at 72 h. Moreover, the degree of DNA fragmentation (expressed as mean channel fluorescence [MCF]) increased with the time of exposure to the toxin. For instance, the MCF for cells exhibiting DNA fragmentation at 24 h was 35; the MCF increased slightly at 48 h to 40. At 72 the MCF increased to 64, indicating that there was extensive DNA fragmentation in these cells. These results are consistent with the notion that Cdt holotoxin-induced DNA fragmentation is both time and dose dependent in lymphocytes; furthermore, the data suggest that DNA fragmentation occurs following G₂ arrest.

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FIG. 4. Kinetics of Cdt-induced DNA degradation. Jurkat cells were treated with Cdt or medium for 24, 48, or 72 h. The cells were then assessed for DNA fragmentation using the TUNEL assay and FACS. The bars indicate regions where there was positive fluorescence; the percentages of cells in these regions that exhibited DNA fragmentation are indicated, and the degrees of fragmentation are indicated by the MCF values. The results are representative of three experiments.

Collectively, our observations are of particular interest because it has previously been shown that Cdt-induced cell cycle arrest culminates in apoptosis (14, 32, 47). Furthermore, several investigators have suggested that Cdt induces cell cycle arrest via DNA degradation; specifically, it has been proposed that the active subunit, CdtB, acts as a DNase (11, 13, 25, 33). Therefore, in the next series of experiments we focused on determining the underlying mechanism that results in increased DNA fragmentation at high Cdt holotoxin concentrations by differentiating between the induction of apoptosis and the direct effects of Cdt on DNA fragmentation. For this purpose we employed two inhibitors of apoptosis, overexpression of the antiapoptotic protein Bcl-2, which inhibits early events associated with the apoptotic cascade, and the caspase-3 inhibitor zvad, which blocks late apoptotic events. We first examined the effects of overexpression of Bcl-2, which has been shown to enhance cell survival in several model systems through inhibition of apoptosis, presumably by preventing development of the MPT (17, 18). For these experiments we employed Jurkat cells transfected with the human bcl-2 gene (Jurkat^bcl-2) and control cells transfected with the empty vector (Jurkat^neo). The protective effects of overexpression of Bcl-2 were first monitored by assessing the development of the MPT. The ability of CdtABC to induce an MPT state was assessed by measuring the _m and generation of reactive oxygen species (ROS). Jurkat cells were treated with medium, a low Cdt dose, or a high CdtABC dose for 18 h and then stained with the fluorescent probes DiOC₆(3) and hydroethidine to measure _m and ROS generation, respectively. Multiparametric FACS analysis indicated that 82.8% (Jurkat^neo) and 90.1% (Jurkat^bcl-2) of the cells exposed to medium alone exhibited bright DiOC₆(3) fluorescence and virtually no ethidium fluorescence [DiOC₆(3)^bright Eth^dim] (Fig. 5, top panel). This is consistent with a high _m and no ROS production in viable cells. Exposure to the low CdtABC dose (0.05 ng/ml) had no effect on DiOC₆(3) fluorescence; 81.8% of the Jurkat^neo cells and 86.2% of the Jurkat^bcl-2 cells exhibited bright DiOC₆(3) fluorescence and no ethidium fluorescence [DiOC₆(3)^bright Eth^dim] (Fig. 5, middle panel). In contrast, exposure to the high dose of CdtABC (5.0 ng/ml) resulted in decreased DiOC₆(3) fluorescence in Jurkat^neo cells [27.7% DiOC₆(3)^dim]. Of the DiOC₆(3)^dim cells, 19.5% also exhibited an increase in ethidium fluorescence (Fig. 5, bottom panel). Thus, the mitochondria of these cells exhibited not only a decline in _m but also increased generation of superoxide anions consistent with development of the MPT. Jurkat^bcl-2 cells were protected from development of the Cdt-induced MPT; 84.3% of these cells still exhibited bright DiOC₆(3) fluorescence (Fig. 5, bottom panel).

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FIG. 5. Cdt induction of MPT in Jurkatneo and Jurkatbcl-2 cells. Control Jurkat cells (Jurkatneo) and cells overexpressing Bcl-2 (Jurkatbcl-2) were treated with Cdt or medium for 18 h. Cells were then stained with DIOC6(3) and hydroethidine (HE) to measure the transmembrane potential and ROS generation, respectively, and were analyzed by flow cytometry. The data are plotted as DIOC6(3) fluorescence versus ethidium fluorescence. The lines indicate the settings for quadrant analysis; the percentages indicate the percentages of cells in each quadrant. The results are representative of three experiments.

We next compared the effects of low and high doses of CdtABC on DNA fragmentation and G₂ arrest in Jurkat^neo and Jurkat^bcl-2 cells (Fig. 6). Small percentages of cells exposed to medium alone exhibited DNA fragmentation (3.7% of Jurkat^neo cells and 2.1% of Jurkat^bcl-2 cells). In the presence of the low CdtABC dose, 9.6% of Jurkat^neo cells exhibited DNA fragmentation, compared to 3.1% of the Jurkat^bcl-2 cells. The percentage of Jurkat^neo cells that exhibited DNA fragmentation in cell cultures treated with the high CdtABC dose was 44.6%; in contrast, DNA fragmentation was observed in only 10.8% of Jurkat^bcl-2 cells. Analysis of the cell cycle indicated that Jurkat^neo cells responded to Cdt holotoxin in a manner similar to that of normal Jurkat cells; the low CdtABC dose induced significant G₂ arrest, while exposure to the high dose did not result in an increased level of G₂ cells but rather resulted in an increase in the level of S-phase cells. In contrast, for Jurkat^bcl-2 cells there was increased G₂ accumulation when the cells were exposed to both low (45.4% G₂) and high (31.3%) CdtABC doses. Thus, overexpression of Bcl-2 not only blocked development of the MPT but also prevented Cdt holotoxin-induced DNA degradation and the accompanying increase in the level of S-phase cells. These results also indicate that prevention of apoptosis does not interfere with Cdt-induced cell cycle arrest.

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FIG. 6. Effect of bcl-2 overexpression on Cdt-induced G2 arrest and DNA fragmentation. Jurkatneo and Jurkatbcl-2 cells were incubated in the presence of medium alone (Ctl), 0.05 ng/ml Cdt, or 5 ng/ml Cdt for 18 h; the cells were then stained with propidium iodide and by TUNEL to measure cell cycle distribution (panels on the right) and DNA fragmentation (panels on the left), respectively. The percentages of cells in the phases of the cell cycle are indicated, as are the percentages of cells exhibiting DNA fragmentation. The results are representative of at least three experiments.

We also employed the caspase inhibitor zvad to block the late stages of apoptosis that ultimately lead to DNA fragmentation associated with this form of cell death. Jurkat cells were exposed to medium or zvad (50 µM) and then treated with the Cdt holotoxin. To confirm that zvad protected cells from late events associated with apoptosis, we first monitored the effects of zvad on Cdt-induced caspase activation. As shown in Fig. 7, cells exposed to medium alone in the presence or absence of zvad exhibited little caspase-3 activation (<1.5%). Exposure to the low CdtABC dose resulted in caspase-3 activation in 26.7% of the cells, compared to caspase activation in 3.5% of the cells pretreated with zvad and then exposed to this dose of Cdt holotoxin. For Jurkat cells treated with the high dose of CdtABC there was a further increase in caspase activation (34.4%); in contrast, zvad reduced the percentage of cells exhibiting caspase-3 activation to 3.8%. DNA fragmentation was observed in 8.8% (low dose) and 58.1% (high dose) of the cells in the absence of zvad (Fig. 8); pretreatment of Jurkat cells with zvad prevented Cdt-induced DNA fragmentation regardless of the dose (<2%). Cell cycle distribution was simultaneously measured in these cells (Fig. 8); in the absence of zvad, treatment of cells with the low dose of CdtABC resulted in 31.4% G₂ cells, and treatment with the high CdtABC dose resulted in 1.8% G₂ cells. For cells exposed to the high concentration 94.9% of the cells were S-phase cells. In contrast, pretreatment with zvad resulted in 37.1% and 45.6% G₂ cells following treatment with the low and high doses of CdtABC, respectively.

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FIG. 7. Inhibition of Cdt-induced caspase activation by zvad. Jurkat cells were pretreated with 50 µM zvad or medium for 30 min; this was followed by addition of medium, 0.05 ng/ml Cdt, or 5 ng/ml Cdt. The cells were then incubated for an additional 18 h and assessed for caspase-3 activation using the fluorescent probe FAM-DEVD-FMK. The regions where there was positive fluorescence are indicated by bars; the percentages of cells exhibiting caspase-3 activation are also indicated. The results are representative of three experiments.

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FIG. 8. Effect of zvad on Cdt-induced G2 arrest and DNA fragmentation. Jurkat cells were pretreated with 50 µM zvad or medium for 30 min; this was followed by addition of either medium (Ctl) or Cdt. Cells were incubated for 18 h and then analyzed for DNA fragmentation (panels on the left) and cell cycle distribution (panels on the right) as described in Materials and Methods. The percentages of cells in the phases of the cell cycle are indicated, as are the percentages of cells exhibiting DNA fragmentation; the regions where there was positive dUTP-FITC fluorescence are indicated by the bars, and the results were based upon control samples.

In a final series of experiments we monitored the affect of Cdt on phosphorylation of the histone H2AX (-H2AX). Phosphorylation of H2AX results from activation of ATM and has been reported to be a useful marker of DNA double-strand breaks (22, 27). As shown in Fig. 9A, control cells (Jurkat^neo) exhibited minimal -H2AX fluorescence (MCF, 10.3); treatment of these cells with 200 pg/ml of Cdt for 4 h (Fig. 9C) resulted in increased phosphorylation of the histone (MCF, 21.1). It should be noted that the amount of -H2AX fluorescence depended on the Cdt dose (data not shown). In contrast, Cdt-induced histone phosphorylation was blocked in Jurkat^bcl-2 cells (Fig. 9B and 9D); untreated Jurkat^bcl-2 cells exhibited an MCF of 7.6, compared with an MCF of 10.4 for cells treated with 200 pg/ml.

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FIG. 9. Effect of Cdt on phosphorylation of H2AX. Jurkatneo (A and C) and Jurkatbcl-2 (B and D) cells were exposed to medium (A and B) or 200 pg/ml Cdt (C and D) for 4 h. The cells were then fixed, permeabilized, and stained with anti-phospho H2AX antibody conjugated to FITC. The results are representative of three experiments; the MCF values are indicated.

Top Abstract Introduction Materials and Methods Results Discussion References

 
There is general agreement that treatment of cells with Cdt holotoxin leads to irreversible G₂ arrest. The time course for cell cycle arrest is cell type specific; in mitogen-activated lymphocytes the peak of arrest is observed 48 to 72 h following exposure to the toxin, and for Jurkat cells the peak of arrest is at 24 h. There is also clear evidence that cell cycle arrest precedes the onset of apoptosis, which occurs in lymphocytes approximately 24 h following G₂ arrest (7, 14, 34, 47). In this study, we observed that at low concentrations (0.00032 to 0.2 ng/ml) Cdt induces a dose-dependent increase in G₂ arrest. However, at concentrations greater than 0.2 ng/ml, the toxin induces a dose-dependent decline in the level of G₂ cells; this decrease correlates with a concomitant increase in the level of S-phase cells. It should be noted that these observations were initially based on a single parametric analysis of cell cycle distribution using propidium iodide; this method relies upon the total DNA content of the cell to determine the cell cycle phase. Further analysis, using BrdU incorporation, failed to identify the cells as S-phase cells. Thus, we ruled out the possibility that the toxin-induced decline in G₂ arrest was the result of transient toxicity and escape from G₂ arrest. We therefore considered other possible causes for these high-dose effects. Since the active toxin subunit, CdtB, has been shown to exhibit DNase activity, we explored the possibility that high doses of Cdt lead to a rapid onset of DNA fragmentation.

The increase in the level of S-phase cells might be related to DNA fragmentation within the arrested G₂ population. DNA fragmentation would be expected to lead to a decline in the total DNA content; this would initially result in cells appearing in the sub-G₂ or S-phase region and eventually in the G₀/G₁ and sub-G0 regions. It is particularly relevant that a routine assay for identifying apoptotic cells monitors the appearance of sub-G₀/G₁ cells (8); this assay is based on the principle that activation of the apoptotic cascade leads to fragmentation and a decrease in the total cellular DNA content. In this context, it was not surprising that we also observed a dose-dependent increase in Cdt-induced DNA fragmentation. Low levels of DNA fragmentation were detected in the presence of CdtABC concentrations that resulted in maximum G₂ arrest; however, the percentage of cells exhibiting DNA fragmentation continued to increase to levels greater than 80% as the concentration of toxin was increased from 0.2 ng to 50 ng/ml. Moreover, dual-parameter flow cytometric analysis indicated that most of the cells initially exhibiting DNA fragmentation at 24 h were indeed in the sub-G₂ region. Further confirmation that DNA fragmentation was the underlying basis for this decline in the G₂ population came from kinetic analysis of both DNA fragmentation and the cell cycle. If DNA fragmentation secondary to cell cycle arrest was responsible for the decline in G₂ cells, it should follow that for cells exposed to low doses of Cdt holotoxin, which lead to G₂ arrest at 24 h, there would eventually be a decline in this population at later times. Indeed, lymphocytes exposed to a low toxin dose resulted in significant G₂ arrest at 24 h; however, at 48 and 72 h, there was a decline in the level of G₂ cells for these cells and there were concomitant increases in the levels of S-phase cells, G₀/G₁, and sub-G₀ cells, respectively. There were also concomitant increases in the percentage of cells exhibiting DNA fragmentation at 48 and 72 h. Likewise, for cells exposed to high doses of Cdt holotoxin, which resulted in a significant increase in "S-phase" cells at 24 h, there was also a further decline in DNA content over time. Thus, these cells began to appear in the sub-G₀ region at 48 h; by 72 h, almost all of the cells were in this region. Furthermore, not only did the percentage of cells exhibiting DNA fragmentation increase, but also the extent of fragmentation in this population increased at 72 h. It should also be noted that at 72 h, the cell cycle distribution of cells exposed to low Cdt doses was similar to that of cells exposed to high doses of Cdt for 48 h. Therefore, it appears that exposure of lymphocytes to Cdt results in not only a dose-dependent increase in the level of G₂ cells but also a dose- and time-dependent increase in DNA fragmentation within the arrested population; exposure to high doses of toxin accelerates this process. It is also interesting that Cortes-Bratti et al. (7) reported similar results for experiments in which they monitored the effects of Cdt on lung fibroblasts over time; these cells first exhibited an increase in the G₂ population at 24 h, and this was followed by a prominent increase in the level of S-phase cells at 48 h.

The observed dose-dependent increase in DNA fragmentation is particularly important with respect to the toxin's mode of action. Several investigators have suggested that the active Cdt subunit, CdtB, has structural homology with DNase I (11, 33). In addition to partial sequence and structural homology, the link between CdtB and DNase I was initially based on two lines of investigation. In one approach the researchers employed mutation of the cdtB gene at loci that are believed to be critical for DNase activity (11, 25, 33). These studies suggested that the same loci are also critical for Cdt function and cell cycle arrest. Other investigators have also demonstrated that Cdt exhibits DNase activity, although CdtB exhibits only 0.01% of the activity of bovine DNase I (10, 11, 29, 33). It should be noted that in these studies the workers assessed the effects of the toxin in terms of its ability to denature or relax supercoiled plasmid DNA. In most of these studies the researchers employed toxin concentrations that were 200- to 20,000-fold greater than the concentration utilized in our study; even under these conditions relatively low levels of DNA fragmentation were observed. In another study, HeLa cells were treated with Cdt and assessed for both cell cycle arrest and the presence of DNA strand breaks (12). Once again, in these studies relatively large doses of Cdt holotoxin and/or CdtB were employed; in some instances CdtB was microinjected (1 µg/cell) into cells (25). While these studies clearly demonstrated the presence of altered DNA, it should be noted that they did not discriminate between direct DNase activity associated with Cdt and the possibility of indirect effects by which toxin treatment led to activation of DNase endogenous to the cell.

It is well documented that activation of the apoptotic cascade leads to activation of endonucleases that results in cellular degradation, including DNA fragmentation. Since observations have clearly demonstrated that treatment of lymphocytes with high concentrations of toxin leads to accelerated DNA fragmentation, we employed two inhibitors of apoptosis in order to establish a relationship between G₂ arrest, activation of the apoptotic cascade, and DNA fragmentation. Overexpression of the antiapoptotic protein Bcl-2 blocks the early stages of apoptosis by interfering with development of the MPT (2, 15, 17, 21, 23). We clearly demonstrated that lymphocytes treated with Cdt holotoxin develop the MPT, which is characterized by a decline in the transmembrane potential [DiOC₆(3) fluorescence] and an increase in ROS generation (ethidium fluorescence). Jurkat^bcl-2 cells which overexpressed Bcl-2 were partially protected from this early apoptotic event. Moreover, Jurkat^bcl-2 cells were resistant to Cdt-induced DNA fragmentation, but they still exhibited susceptibility to Cdt-induced G₂ arrest. It was particularly relevant that the high-dose effects of toxin, which led to decreases in the level of G₂ cells (and increases in the level of S-phase cells) in control Jurkat cells, were less pronounced in the Jurkat^bcl-2 cells. Likewise, the caspase inhibitor zvad has been shown to inhibit late apoptotic events, such as caspase-3 activation and DNA fragmentation. Cdt-treated Jurkat cells exhibited caspase-3 activation; however, in the presence of zvad, this apoptotic event was blocked. As observed with bcl-2 overexpression, cells treated with zvad did not exhibit DNA fragmentation in the presence of high doses of Cdt; nonetheless, both low and high doses of toxin induced G₂ arrest in these cells. Thus, we concluded that the extensive DNA fragmentation that we and other workers have observed in cells treated with high concentrations of toxin results from activation of the apoptotic cascade and is secondary to G₂ arrest.

Consistent with this interpretation are our observations regarding phosphorylation of the histone H2AX. Phosphorylation of this histone has been shown to be an early indicator of both DNA damage associated with double-strand breaks and activation of the apoptotic cascade (18, 22, 27). Indeed, it has been shown previously that Cdt-treated cells exhibit ATM-dependent phosphorylation of H2AX (49); these results were interpreted as evidence that Cdt functions as a DNase and induces DNA damage. Therefore, we investigated whether activation of this nuclear sensor of DNA damage was directly due to Cdt exposure or due to activation of the apoptotic cascade. Our results indicate that overexpression of Bcl-2 blocks not only apoptosis and DNA fragmentation but also phosphorylation of H2AX. Thus, these results suggest that Cdt-induced DNA damage and the resulting activation of the ATM pathway may in fact result from indirect effects of the toxin and be a direct consequence of activation of the apoptotic cascade.

In summary, we propose that exposure of lymphocytes to Cdt first results in G₂ arrest by mechanisms that are most likely related to activation of the G₂ checkpoint (7, 9, 42, 46). Cdt-induced cell cycle arrest is irreversible and eventually leads to activation of the apoptotic cascade. Our observations also indicate that the cell death pathway involves development of the MPT and eventually caspase-3 activation. Based upon this sequence, one would not expect nonproliferating cells to become apoptotic in the presence of Cdt. Indeed, we have previously shown that Cdt does not induce apoptosis in primary lymphocytes unless they are activated by mitogen (47). It is also clear from the present study that high doses of Cdt lead to more rapid onset of the apoptotic cascade. While the mechanism responsible for this effect is not known, it is unlikely to be due to the direct effects of CdtB functioning as a DNase. Our study supports the notion that the vast majority of DNA fragmentation that we and other workers have reported in association with exposure to high concentrations of Cdt and possibly the DNA fragmentation resulting from exposure to low doses as well are the result of activation of the apoptotic cascade (14, 34, 35, 47). These observations are consistent with those of Sert et al. (42), who demonstrated that Cdt-induced G₂ arrest is independent of DNA damage. Thus, while there have been a number of reports that support the hypothesis that DNA degradation is involved in the mechanism of action of Cdt, these studies did not exclude the possibility that the effects were mediated indirectly by endogenous enzymes. In fact, in light of the extremely low doses of Cdt required to induce G₂ arrest (picograms) in lymphocytes and the relatively high toxin concentrations employed by other investigators to induce DNA fragmentation (micrograms), it is not clear that the primary Cdt mode of action is mediated directly by DNA fragmentation. Nonetheless, it should be emphasized that our observations do not rule out the possibility that a component of the Cdt mode of action is indeed related to low-level DNA damage. It is evident from these observations that the mechanism by which Cdt intoxicates cells and induces irreversible cell cycle arrest is likely to be more complex than the mechanism originally proposed.

 
This work was supported by Public Health Service grant DE06014 from the National Institute of Dental and Craniofacial Research.

 
* Corresponding author. Mailing address: Department of Pathology, University of Pennsylvania School of Dental Medicine, 240 South 40th Street, Philadelphia, PA 19104-2030. Phone: (215) 898-5959. Fax: (215) 573-2050. E-mail: shenker{at}pobox.upenn.edu.

Editor: J. T. Barbieri

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Infection and Immunity, April 2006, p. 2080-2092, Vol. 74, No. 4
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