DNA damage-induced [Zn2+]i transients: correlation with cell cycle arrest and apoptosis in lymphoma cells

Paul J. Smith1, Marie Wiltshire1, Sharon Davies1, Suet-Feung Chin2, Anthony K. Campbell3, and Rachel J. Errington3

Departments of 1 Pathology and 3 Medical Biochemistry, University of Wales College of Medicine, Cardiff, CF14 4XN; and 2 Cambridge Institute for Medical Research, Addenbrooke's Hospital, Cambridge CB2 2QH, United Kingdom


    ABSTRACT
TOP
ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
REFERENCES

Reactive changes in free intracellular zinc cation concentration ([Zn2+]i) were monitored, using the fluorescent probe Zinquin, in human lymphoma cells exposed to the DNA-damaging agent VP-16. Two-photon excitation microscopy showed that Zinquin-Zn2+ forms complexes in cytoplasmic vesicles. [Zn2+]i increased in both p53wt (wild type) and p53mut (mutant) cells after exposure to low drug doses. In p53mut cells noncompetent for DNA damage-induced apoptosis, elevated [Zn2+]i was maintained at higher drug doses, unlike competent p53wt cells that showed a collapse of the transient before apoptosis. In p53wt cells, the [Zn2+]i rise paralleled an increase in p53 and bax-to-bcl-2 ratio but preceded an increase in p21WAF1, active cell cycle arrest in G2, or nuclear fragmentation. Reducing [Zn2+]i, using N,N,N',N'-tetrakis(2-pyridylmethyl)ethylenediamine, caused rapid apoptosis in both p53wt and p53mut cells, although cotreatment with VP-16 exacerbated apoptosis only in p53wt cells. This may reflect changed thresholds for proapoptotic caspase-3 activation in competent cells. We conclude that the DNA damage-induced transient is p53-independent up to a damage threshold, beyond which competent cells reduce [Zn2+]i before apoptosis. Early stress responses in p53wt cells take place in an environment of enhanced Zn2+ availability.

flow cytometry; two-photon laser scanning microscopy; zinc


    INTRODUCTION
TOP
ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
REFERENCES

THE TRANSITION METAL ZINC is a physiologically important divalent cation in biological systems (43). The availability of Zn2+ can affect various macromolecular processes, including DNA synthesis, microtubule polymerization, apoptosis, and gene expression (5, 10). The role of Zn2+ in protein structure and function (12, 13) is highlighted by the frequent participation of zinc-sulfur cluster-containing proteins in replication, recombination, and transcription, with the zinc finger structural domain being an important DNA-binding motif. In most cells the free intracellular concentrations of Zn2+ ([Zn2+]i) are extremely low, with the majority being bound to proteins (25, 30, 36) at various subcellular locations (20). Such locations provide potentially labile stores of Zn2+ for mobilization. Metallothioneins (MTs) are the major intracellular Zn2+-binding proteins, and changes in their expression will impact on free Zn2+ levels and, consequently, cation availability for cellular processes. For example, the ability of downregulation of MT to induce growth arrest and apoptosis in human breast carcinoma cells (1) may be effected through changes in Zn2+ pools.

MT expression can change in response to DNA damage in vivo (3, 22) and appears to be integrated into the stress responses of cells (6, 14). Thus there is a rationale for expecting shifts in free Zn2+ levels to occur in cells undergoing DNA damage-induced stress responses including caspase activation and the triggering of apoptosis (10, 18). The control of apoptosis is an essential process in the development and homeostasis of all metazoans. One level of control is through the inhibitor-of-apoptosis proteins (IAPs) that suppress cell death by inhibiting the activity of caspases; this inhibition is performed by Zn2+-binding domains of IAPs (48). A recent study (17) has indicated that stress response genes p53, gadd45, and c-fos as well as caspase-3 activity appeared to be modulated by cellular Zn2+ status. Growing normal human bronchial epithelial cells in Zn2+-deficient medium resulted in elevation of p53 mRNA, nuclear p53 protein, and gadd45 mRNA, and caspase-3 activity was marginally reduced (17). The active and acute depletion of intracellular Zn2+, using the chelator N,N,N',N'-tetrakis(2-pyridylmethyl)ethylenediamine (TPEN), can induce protein synthesis-dependent neuronal apoptosis in mouse cortical cell cultures (29) or cell death in neuro-2A cells (33) and promotes the formation of internucleosomal DNA fragments in peripheral blood lymphocytes damaged by hydrogen peroxide (44). TPEN has also been found to induce apoptosis in mouse neuroblastoma cells, and this process could be prevented by equimolar exogenous Zn2+ (35). Such observations suggest that reductions of intracellular free Zn2+ alone can result in both cellular stress and the triggering of apoptosis and may act in concert with DNA damage to enhance DNA fragmentation. Here we have sought to understand the sequence of any DNA damage-induced shifts in free Zn2+ in cell lines with the differential ability to mount a p53 stress response and, consequently, the differential capacity to engage the apoptotic pathway. Our aim was to reveal the free Zn2+ environment that exists during the mounting of p53-driven stress responses and how this may change in cells undergoing suprathreshold levels of DNA damage and progression to apoptosis.

The tumor suppressor TP53 gene plays a central role in the cellular response to DNA damage. TP53 is the most frequently mutated gene in human cancer and encodes a Zn2+-requiring transcription factor that can control the traverse of the cell cycle and the commitment to apoptosis of cells exposed to DNA-damaging agents (38). In cells with functional p53-dependent pathways, DNA damage triggers both G1 and G2 arrest of cells (15, 31). Dysfunction of p53-dependent pathways can allow tumor cells to evade cell cycle checkpoint controls and apoptosis (32). Cells lacking p53 or its downstream effector, p21WAF1, fail to maintain a G2 arrest following gamma irradiation (8, 47). The complex tertiary structure of p53 incorporates a Zn2+-stabilized DNA-binding domain, and mutations affecting this domain of the protein appear to be associated with significantly shorter cancer-related survival for colorectal cancer disease (7). It is possible that changes in cellular Zn2+ levels can modulate p53 function. Attempts to manipulate protein conformation and function by the removal of Zn2+ have revealed the generation of modified conformational forms of p53 with decreased DNA-binding activity (45), whereas protein refolding results in increased DNA binding, expression of the downstream effector p21WAF1, and cell cycle delay (45). Thus metalloregulation, comprising intracellular Zn2+ fluxes, could affect p53-dependent driven responses including cell cycle delay and cell death (10).

The continuous noninvasive tracking of free Zn2+ in single cells, over the time courses required to identify the initiation of cell cycle arrest and cell death, is problematic and not currently possible. Here we have opted to identify DNA damage-induced changes in the low levels of free Zn2+ in individual cells and to correlate these events, using flow cytometry, with the stress responses of p53 stabilization, cell cycle arrest, and apoptosis. The probe Zinquin [(2-methyl-8-p-toluenesulfonamido-6-quinolyloxy)acetic acid] has been used to detect free Zn2+ in intact cells (50, 51). Intact cells, treated with Zinquin ester (Zinquin E), can be loaded with the probe through intracellular esterase conversion. Zinquin can be used, in a manner analogous to that of the nonratiometric probes for free Ca2+, to detect free or loosely bound (labile) intracellular Zn2+ by one-photon (4) or two-photon excitation (37) and fluorescence analysis. In the current study we have used continuous exposure to the DNA-damaging topoisomerase II inhibitor VP-16 (etoposide) to induce cycle delay and cell death in lymphoma cells. Under such exposure conditions (39, 41), VP-16 progressively induces DNA damage through the trapping of topoisomerase II enzyme molecules on DNA in a form that leads to strand break formation and the progressive generation of stress signals for arrest in G2 and for the triggering of cell death. The results reveal early increases in the low levels of free Zn2+ in cells undergoing DNA damage induction irrespective of p53 function, whereas a reduction in free Zn2+ occurs in cells engaging apoptosis. The results show differences between the engagement of apoptosis through Zn2+ chelation and DNA damage but indicate that the two inducers can cooperate in the promotion of cell death.


    MATERIALS AND METHODS
TOP
ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
REFERENCES

Cell Lines and Culture Conditions

Two human follicular B lymphoma cell lines were used in this study. DoHH2 cells were a kind gift from Dr. J. C. Kluin-Nelemans (Leiden, The Netherlands) (21), and SU-DHL-4 cells were provided by Dr. F. E. Cotter (16). DoHH2 was routinely maintained in RPMI 1640 supplemented with 5% FCS. SU-DHL-4 cells were cultured in RPMI 1640 supplemented with 10% FCS (Autogen Bioclear). HL-60 cells (ATCC CCL-240; acute promyelocytic leukemia) were included in this study as an immunoblotting control for p21WAF1/SDI1/CIP1 and were grown in RPMI 1640 supplemented with 20% FCS. All growth media were supplemented with 100 U/ml penicillin, 100 µg/ml streptomycin, and 2 mM glutamine. The cell lines were passaged twice weekly at an initiating density of 5 × 104 cells/ml cultured at 37°C in a humid atmosphere of 5% CO2-95% air. The p53 status of the DoHH2 (wild type) and SU-DHL-4 (mutant) cell lines was confirmed by immunocytochemistry, immunoblotting, and analysis of exons 5-8 SSCP (P. J. Smith, unpublished data).

Reagents

Zinquin E {[2-methyl-8-(4-methylphenylsulfonylamino)quinolinyl]oxyacetic acid ethyl ester} was purchased from Alexis and stored as a 5 mM stock solution in ethanol at 4°C. VP-16 (VP-16-213; VEPESID) was provided as a 34 mM stock solution (Bristol Meyers Pharmaceuticals, Syracuse, NY) and stored at 4°C. Fluorescein-conjugated annexin V (annexin V-FITC) was purchased from Pharmingen (Becton Dickinson UK, Oxford, UK.). Propidium iodide (PI) was obtained as a 1 mg/ml solution in H2O (Molecular Probes Europe, Leiden, The Netherlands).

Flow Cytometric Analysis of p53 Content

Approximately 1-5 × 106 cells were pelleted by centrifugation at 1,250 g for 10 min at 4°C. The cell pellet was loosened by gentle tapping and then incubated for 15 min on ice with 0.25% paraformaldehyde (PFA) made up in PBS. The cells were pelleted and washed once with PBS. The cell pellet was washed once with PBS and fixed with 5 ml of 70% ethanol at -20°C for 2 h or overnight. The fixed cells were pelleted by centrifugation and rehydrated by being washed in PBS. The cells were permeabilized with 0.25% Triton X-100 in PBS for 5 min on ice and rinsed with PBS. The cells were then incubated with 100 µl of primary antibody [Pab-6 (clone DO-1), anti-human pantropic p53 mouse IgG2a antibody; Oncogene Sciences] diluted 1:100 in antibody dilution buffer (PBS, 5% nonfat milk, and 0.1% Triton X-100) for 1 h at room temperature or overnight at 4°C. The cells were rinsed with PBS and incubated with FITC-conjugated secondary antibody diluted in antibody dilution buffer for 1 h at room temperature. Cells were rinsed with PBS and resuspended with 1 ml of 5 µg/ml PI in PBS and 0.1% RNase A.

Immunoblotting

Approximately 2 × 106 cells were pelleted by centrifugation at 1,250 g for 5 min, washed once with ice-cold PBS, and lysed with 100 µl of sample loading buffer (34). Cell lysates were boiled for 5 min to denature proteins and were transferred immediately onto ice. Cellular debris and DNA material were removed by centrifugation, and lysates were electrophoresed on 12% SDS-polyacrylamide gels (24). Proteins were electrophoretically transferred to nitrocellulose membranes (Immobilon-P; Millipore) in transfer buffer (25 mM Tris, pH 8.3, 192 mM glycine, and 20% methanol) (42), and gels were stained to check for equal loading. Membranes were incubated in 5% skim milk made up in wash buffer (100 mM Tris, pH 7.5, 0.9% wt/vol NaCl, and 0.1% Tween 20) at room temperature for 1 h to minimize nonspecific binding of antibody before washing and antibody incubation. Antibodies used were diluted in wash buffer and comprised anti-human p21WAF1/SDI1/CIP1 (clone 6B6, mouse IgG1; Pharmingen) used at 1:50 dilution, anti-human bcl-2 (mouse IgG; kindly supplied by Dr. F. E. Cotter) used at 1:50 dilution, and anti-human bax (rabbit IgG; Santa Cruz) used at 1:50. The membranes were washed and incubated with appropriate dilutions (1:2,000-1:5,000) of horseradish peroxidase-linked secondary antibody (Amersham) at room temperature for 1 h. Immune complexes were detected by enhanced chemiluminescence detection (ECL kit; Amersham Biosciences, Amersham, UK).

Analysis of Cell Cycle Changes and Apoptosis

Cell cycle analysis and laser light scatter. Cell cycle analysis as Triton-X-permeabilized and ethidium bromide-stained nuclei was performed as described previously (40). The analysis of laser side/90° light scatter was used to identify nuclear fragments and abnormal nuclear structure.

In situ terminal deoxynucleotidyl transferase assay. Approximately 1 × 107 cells/ml (untreated or treated with drugs) were pelleted by centrifugation at 1,250 g for 10 min at 4°C. The pellets were washed once with PBS and fixed in suspension with 5 ml of 4% PFA in PBS at room temperature with gentle agitation for 15 min. The fixed cells were rinsed with PBS, and pellets were resuspended with PBS to obtain 1 × 107 cells/ml. Cell suspension (10 ml) was aliquoted onto each well on 15-well poly-lysine-treated slides and incubated for 6-18 h in a humidified chamber to allow cell attachment. Slides were washed in PBS followed by H2O and then air dried. The cells in each well were incubated with 10 µl of direct reaction mixture [78 µl dH2O, 20 µl of 5× terminal deoxynucleotidyl transferase (TdT) buffer, 1 µl of TdT enzyme, and 1 µl of fluorescein 11-dUTP (Fluorogreen)] for 1 h at 37°C in a humidified chamber. Slides were rinsed with PBS and H2O. Cells were counterstained with PI (0.04 µg/ml in PBS) for 2 min at room temperature, rinsed, and mounted with VectorShield. Samples were examined by using a Bio-Rad MRC-600 confocal laser scanning microscope operating in dual-channel fluorescence mode for the simultaneous imaging of PI vs. TdT positivity. Cells that had intensely stained nuclear bodies (PI; red) with positive TdT staining (green) were counted (n > 100) as apoptotic cells containing single-strand DNA breaks, whereas normal cells were those with diffuse nuclear staining and negative staining for TdT.

Annexin V binding. Samples were prepared (46) for the detection of annexin V-FITC surface binding to cells undergoing apoptotic changes and costained with PI to detect loss of plasma membrane integrity.

Loading Conditions for Zn2+ and Zinquin

Cells were grown in asynchronous culture to ~6 × 105 cells/ml, resuspended in loading buffer [Hanks' balanced salt solution lacking Ca2+ and Mg2+ supplemented with 20 mM HEPES (pH 7.4)], and held at 37°C. Cells were then exposed to 0-100 µM Zn2+ with or without sodium pyrithione (1 µM) and incubated for 30 min at 37°C. After Zn2+ or control loadings, cells were washed twice rapidly in loading buffer (centrifugation for 30 s at 6,500 rpm with a MicroCentaur microfuge) and resuspended at 6 × 105 cells/ml. Suspensions were sham treated or exposed to 25 µM Zinquin E and incubated for 30 min at 37°C, and 1 µg/ml PI was present for the final 10-min period of incubation. Cell suspensions were analyzed directly by flow cytometry or mounted on microscope slides for imaging.

Analysis of Zinquin-Zn2+ in Single Cells

Flow cytometry. After experimental manipulations were performed, cells were analyzed immediately by using a fluorescence-activated cell sorting Vantage flow cytometer (Becton Dickinson Immunocytometry Systems, San Jose, CA) incorporating an Innova Enterprise II argon ion laser (Coherent, Santa Clara, CA) emitting 488-nm and multiline ultraviolet (MLUV; 351-355 nm, 30 mW) wavelengths. Forward scatter (FSC; master signal) and side scatter (SSC) were acquired in linear mode for 10,000 cells. PI (cell integrity probe) and ethidium bromide (DNA stain for cell cycle analysis) were excited at 488 nm, and fluorescence signals were detected with a 585/42-nm band-pass filter. Zinquin fluorescence (350- to 360-nm excitation wavelength range, 485-nm emission wavelength maximum) originating from the MLUV laser excitation was collected in linear mode at a photomultiplier position protected by DF 424/44-nm and 510-nm dichroic filters. CELLQuest software (Becton Dickinson Immunocytometry Systems) was used for signal acquisition and analysis. Data are expressed as mean fluorescence intensity for populations of single cells. CELLQuest software was used to generate the Kolmogorov-Smirnov statistical analyses for two-sample tests for histograms. The K-S analysis tests whether two selected histograms are different. The calculation computes the summation of the curves and finds the greatest difference between the curves, and then, assuming that the two selected histograms are from the same population, the analysis tests the probability of that difference (D) being as large as measured (49).

Two-photon excitation laser scanning microscopy. Cells were treated and loaded with Zinquin E. These preparations were placed onto Labtek chambered cover glass (NUNC catalog no. 178565) and mounted onto an inverted microscope (Zeiss Axiovert 100). Three-dimensional (3-D) (x, y, z) images were acquired through the cells by using a laser scanning microscope with two-photon excitation (Bio-Rad 1024MP; Bio-Rad Microscience, Hemel Hempstead, UK). Two-photon excitation (by which infrared light can be used to elicit fluorescence from a UV-excitable fluorochrome) was achieved by using a mode-locked 10-fs pulsed Titanium-Sapphire laser (Verdi-Mira 900; Coherent Lasers, Cambridge, UK) tuned to 780 nm, and fluorescence emission was acquired between 460 and 650 nm. All experiments were conducted with the use of a ×60, 1.4-NA, oil-immersion objective lens. For visualization, the entire 3-D image was projected, with the use of a maximum intensity algorithm, into a single two-dimensional view by using standard analysis software (LaserSharp v3.0; Bio-Rad Microscience).


    RESULTS
TOP
ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
REFERENCES

Apoptosis Induced by DNA Damage or Zn2+ chelation

Figure 1, A-D, shows typical results for the measurement of apoptosis induction and progression to cell death in DoHH2 cell populations exposed to VP-16, Zinquin E, or TPEN. The lower right quadrants in Fig. 1, A-D, delineate cells with high annexin V binding but with intact plasma membranes, defining apoptosis without loss of cellular integrity. After a short period of exposure (<= 4 h) to either Zinquin E or VP-16, cell populations produced profiles the same as those for control cultures with a low background of preexisting apoptotic cells. Figure 1E shows that continued exposure (>4 h) to VP-16 resulted in a progression of DoHH2 cells to apoptosis (lower right quadrant characteristics) followed by loss of membrane integrity (upper right quadrant characteristics). This finding contrasts with the marked resistance of the SU-DHL-4 cells to VP-16-induced apoptosis (Fig. 1F). Treatment of DoHH2 cells for 4 h with TPEN resulted in a clear increase in apoptotic fraction (Fig. 1B, lower right quadrant) that retained plasma membrane integrity (no increase in upper right quadrant), suggesting a rapid commitment to cell death upon Zn2+ chelation.


View larger version (34K):
[in this window]
[in a new window]
 
Fig. 1.   Kinetics of apoptotic commitment and cell death in populations of DoHH2 cells monitored by flow cytometry. A-D show typical dot plots for annexin V binding vs. cellular integrity [propidium iodide (PI) exclusion] with the quadrant analysis regions for intact viable cells (lower left), apoptotic cells (lower right), nonviable cells with disrupted plasma membranes (upper right), and cellular debris (upper left). A: control. B: exposure to 25 µM N,N,N',N'-tetrakis(2-pyridylmethyl)ethylenediamine (TPEN) for 4 h. C: exposure to 25 µM Zinquin E for 4 h. D: exposure to 2.5 µM VP-16 for 4 h. E and F show time-dependent changes in DoHH2 and SU-DHL-4 cells in the frequency of apoptotic (open circle ) and nonviable () cells during exposure to 2.5 µM VP-16. In E and F, the dashed and solid horizontal lines indicate the range of control values for apoptotic and nonviable cells, respectively.

DNA Damage-Induced Cell Cycle Arrest

p53-independent accumulation in G2 provides a means of evaluating the DNA damage-associated stress responses of the two cell lines. The VP-16 dose dependency (Fig. 2, A and B) and kinetics (Fig. 2, C and D) of cell cycle arrest in the two lymphoma cell lines were determined by analyzing all cells with normal range DNA content. In general the two cell lines showed similar patterns of response, in keeping with their similar cell cycle times, upon long-term (24 h) exposure to 0.125-2 µM VP-16, with DoHH2 cells (Fig. 2A) displaying a reduced extent of cell cycle redistribution compared with SU-DHL-4 cells (Fig. 2B). SU-DHL-4 cells showed a dose-dependent increase in G2/M arrest peaking at 0.5 µM, at which point G1 emptying is complete. At doses >0.5 µM VP-16, an increasing number of cells also became trapped in S phase. These data indicate the lack of any significant G1/S arrest in SU-DHL-4 cells. The accumulation of DoHH2 cells in G2/M was up to threefold less than that for SU-DHL-4 cells at 0.5 µM VP-16 with evidence of the retention of cells in G1.


View larger version (30K):
[in this window]
[in a new window]
 
Fig. 2.   Drug-induced cell cycle arrest in DoHH2 (A and C, open symbols) and SU-DHL-4 cultures (B and D, filled symbols). A and B show the dose dependency of cell cycle perturbations after 24-h exposure to VP-16. C and D show the time course of changes in cell cycle distribution following treatment with 0.5 µM VP-16. Horizontal lines represent the maximum range of values for G1 (dashed lines) and G2/M (dotted lines) for parallel untreated control cultures. Data are means ± SD for 3 experiments. Circles, G1 phase; triangles, S phase; squares, G2/M phase.

Following the kinetics of arrest at the 0.5 µM VP-16 level (Fig. 2, C and D) revealed similar patterns of arrest/delay, with redistribution outside the normal range becoming apparent at >= 4 h of drug exposure. The kinetics also revealed that the two cell lines show similar rates of exit from G1, despite initial differences in the G1 fractions of untreated cells, over the initial 8-h exposure period and a progressive accumulation of cells in G2/M up to 20 h of drug exposure. Cell cycle checkpoint reversal experiments using caffeine indicated that SU-DHL-4 cells arrested in G2/M can be forced to progress through to G1 with a concomitant loss of their elevated cyclin B1 levels (S.-F. Chin and P. J. Smith, unpublished data). This finding suggests that VP-16-treated SU-DHL-4 cells exhibit a sustained G2 delay under continuous VP-16 exposure conditions.

Taken together, these data reveal that exposure to 0.5 µM VP-16 is a critical threshold below which the initial cell cycle changes are comparable in the two cell lines, whereas beyond that level of additional stress factors, such as apoptotic engagement in DoHH2, affect distribution patterns. The data provide a VP-16-induced stress profile that permits a rational search for linked changes in free [Zn2+]i.

Early Changes in Cell Cycle/Apoptosis Regulators in DoHH2 Cells During VP-16 Treatment

To determine whether VP-16 treatments could differentially engage apoptosis to the level of DNA fragmentation/DNA strand break generation, we examined the frequency of cells showing DNA fragmentation after 24 h of drug exposure. Figure 3A demonstrates that the majority of DoHH2 cells treated with >= 0.25 µM VP-16 are destined to become apoptotic. The SU-DHL-4 cell line showed resistance to fragmentation. To link any changes in the expression of p53 in DoHH2 cells with the engagement of the DNA fragmentation process, we used differential stabilization of nuclear bodies by ethanol alone, in which nuclear integrity is fragile, compared with a cross-linking procedure in which nuclear integrity is better preserved by pretreatment with PFA (see MATERIALS AND METHODS). Figure 3B shows the low levels of p53 detected in control DoHH2 cells. VP-16-treated and PFA-prefixed cells showed high levels of p53 induction in all phases, but particularly in the G2/M arrested population. Without PFA prefixing, the high p53 expression events were lost to a sub-G1 fraction (Fig. 3B, arrow), indicating that their nuclear bodies have undergone disruption. Thus, even at 12 h of drug treatment, there is clear evidence that p53 expression has not been induced in all cells and that these cells preserve nuclear integrity and would contribute to the progressive arrest shown in Fig. 2C.


View larger version (44K):
[in this window]
[in a new window]
 
Fig. 3.   Induction of apoptosis and changes in p53, bcl-2, and bax proteins in VP-16 treated human cells. A: VP-16 induction of DNA fragmentation/apoptosis as determined by an in situ TdT assay for DNA fragmentation. Data are means ± SD for 3 experiments. open circle , DoHH2 cells; triangle , SU-DHL-4 cells. B: effect of fixation conditions on the flow cytometric determination of cellular p53 content in control and VP-16 treated DoHH2 cells. Paraformaldehyde (PFA) plus ethanol fixation preserves cell integrity for the quantification of p53 levels vs. the indicated cell cycle position. Ethanol fixation alone results in a loss of p53-positive events to the sub-G1 (fragment) DNA region in drug-treated cultures (arrow). The bold line indicates the maximum level (upper limit for >= 95% of events) of fluorescence detected in untreated controls. C: flow cytometric determination of changes in p53 content of PFA plus ethanol-fixed cells as a function of cell cycle age and duration of exposure to VP-16 (0.5 µM). The bold line is reproduced from B. The data show early increases in all cell cycle stages with persistence of elevated levels as cells progress to arrest in G2/M. D: changes in p21WAF1, bcl-2, and bax content of DoHH2 cells with duration of exposure to VP-16 (0.5 µM) detected by immunoblotting. Whole cell extracts of SU-DHL-4 (p53 mutant) and phorbol ester-treated HL-60 cells (HL-60+TPA) are included as marker controls.

Figure 3C shows that the time course of p53 changes for PFA-prefixed DoHH2 cells. Many cells (>50% population), in all phases of the cell cycle, showed an early (4 h) increase in p53 levels (beyond the control levels shown by the bold bar and defined in Fig. 3B), with the remainder showing p53 levels within the range of the untreated control. This elevation of p53 was maintained in cells as they progressed through to G2/M arrest, with the cells remaining in G1 and S phase showing progressive loss of an elevated p53-specific signal. The presence of apoptotic internucleosomal DNA cleavage was confirmed by gel electrophoresis (data not shown). Parallel studies using pulsed-field electrophoresis to detect the initial high-molecular-weight fragmentation of DNA indicated that cleavage is not detectable for 0.5 µM VP-16-treated DoHH2 cells before 3 h of drug exposure (S.-F. Chin and P. J. Smith, unpublished data).

Figure 3D shows the parallel analysis of changes in p21WAF1, bcl-2, and bax levels upon drug exposure as indicators of cellular stress responses (23). The results indicate the early activation of p21WAF1 by 3 h of drug exposure and its greater expression during cycle arrest with an apparent progressive increase in the bax-to-bcl-2 ratio evident after 2 h of drug exposure.

Thus the VP-16 response study identifies treatment conditions under which cycle arrest will ensue in both cell lines with similar kinetics while apoptosis can be differentially engaged. The continuous presence of the DNA-damaging agent VP-16 appears to sustain the elevations in p53 and p21WAF1 that precede the cell cycle perturbations in DoHH2. p53 (wild type)-dependent changes in gene expression were not observed in SU-DHL-4 cells. The flow cytometric analysis demonstrated that DoHH2 cells that become arrested in G2/M do so with a sustained increase in p53 and commitment to apoptotic DNA fragmentation.

Subcellular Location of Free Intracellular Zn2+ Revealed by the Zinquin E Probe

To enable interpretation of the flow cytometric studies of total cellular fluorescence, we used two-photon microscopy (37) to detect the subcellular location and intensity of intracellular Zinquin fluorescence. We examined cells expressing basal levels of free intracellular Zn2+ and cells preloaded with extracellular Zn2+ [i.e., using the ionophore sodium pyrithione as described previously (50)]. Two-photon excitation laser scanning microscopy of Zinquin complexes within these B-cell lymphomas revealed that fluorescence was enhanced by extracellular Zn2+ and was punctate, being localized into a subcellular compartment excluded from the nucleus.

Figure 4, a-e, shows the distribution of Zinquin-Zn2+ complexes in SU-DHL-4 cells and the effects of ionophore loading with Zn2+. The images indicate that the basal level of fluorescence associated with Zinquin E incubation is very low compared with that achieved through sodium pyrithione-mediated preloading with Zn2+, although the punctate patterns appear to be similar. Preincubation of cells with Zn2+ without ionophore suggests that cells can accumulate and sequester Zn2+ in stores accessible to Zinquin. Colocalization studies using the novel cell-permeant DNA dye DRAQ5 (37) indicated that the properties of Zinquin preclude its ability to locate or remain within the nuclear compartment or that ionophore loading of Zn2+ results in metal cation sequestration within nonnuclear vesicles (images not shown). We also observed that mitotic cells lacking a nuclear membrane and cells with clearly defined apoptotic nuclear morphologies also show punctate fluorescence of Zinquin-Zn2+ complexes (images not shown).


View larger version (63K):
[in this window]
[in a new window]
 
Fig. 4.   Distribution of Zinquin-Zn2+ complexes in SU-DHL-4 cells. Suspensions of cells were pretreated with or without 25 µM intracellular Zn2+ and with or without 1 µM sodium pyrithione, then incubated with Zinquin E before being placed in a microscope observation chamber. Images show representative views of 2 cells per condition. Two-photon excitation was used to excite samples at a 780-nm wavelength, and all fluorescence was detected above 470 nm. Optical sections were acquired through the cells and projected maximally into a single view: a, autofluorescence of untreated cells; b, cells loaded with Zinquin E alone; c, cells pretreated with Zn2+ alone; d, cells pretreated with sodium pyrithione alone; and e, cells pretreated with both Zn2+ and sodium pyrithione.

Subpopulation Analysis of Early Changes in [Zn2+]i During VP-16 Exposure

Because our aim was to detect shifts in free [Zn2+]i in these complex cell populations responding to a cytotoxic agent, it was clearly important to exclude dead and/or dying cells from the analyses. Imaging indicated that although such cells esterase-convert Zinquin E, their compromised membrane integrity could result in the reporting of anomalous free [Zn2+]i (data not shown). Thus viability was defined in terms of membrane integrity and was determined by PI exclusion (see Fig. 1, A and C). Preliminary experiments (data not shown) confirmed that flow cytometry could detect the extent of ionophore-mediated loading of extracellular Zn2+ and established standard conditions for Zinquin E loading.

To investigate DNA damage-induced changes in [Zn2+]i, our strategy was to investigate the early shifts in Zinquin fluorescence for VP-16 exposures capable of initiating early molecular events (i.e., <4 h) that precede cycle arrest or have the potential to fully engage apoptosis. With the use of short drug incubation periods, reactive changes in [Zn2+]i could be analyzed without the complication of longer term cell cycle redistribution. This analysis period also corresponds to the period during which an enforced fall in free [Zn2+]i can induce apoptosis in DoHH2 cells (Fig. 1B) and during which VP-16 can induce significant changes in cell cycle/apoptosis regulators (Fig. 3, C and D) without the immediate expression of apoptotic changes per se (Fig. 1E).

In all experiments in which control cells were treated with Zinquin E alone, with cellular fluorescence reflecting basal levels of free [Zn2+]i, we observed a subpopulation of PI-negative cells with high levels of Zinquin fluorescence (Fig. 5A). The dot plot also shows that the normal background of PI-positive dead DoHH2 cells shows increased Zinquin fluorescence. The increasing Zinquin fluorescence positivity in R2 is not associated with an increase in PI signal despite the assay sensitivity (note the logarithmic scale). It is important to note that the Zinquin and PI signals are generated through excitation at spatially separated laser beams, and therefore the high-[Zn2+]i subpopulation is not an artifact of coexcitation. To define this population of elevated [Zn2+]i cells, we arbitrarily set lower (>2-fold control population mean) and upper (<8-fold control population mean) limits for Zinquin fluorescence. Accordingly, the mean percentages of the elevated [Zn2+]i subpopulations of PI-negative cells in control cultures were 6.72 ± 4.45 and 7.22 ± 5.26% for DoHH2 and SU-DHL-4 cells, respectively.


View larger version (35K):
[in this window]
[in a new window]
 
Fig. 5.   Typical flow cytometric dot plots showing Zinquin fluorescence (FL5-H parameter) vs. cellular integrity (PI exclusion; FL2 parameter) in populations of control (A) and drug-treated (B; 0.25 µM VP-16 for 2.5 h) DoHH2 cells. Intact PI-negative cells show a subpopulation with up to 4-fold elevated [Zn2+]i. R1, low-[Zn2+]i cells; R2, high-[Zn2+]i cells. The dot plots indicate the anomalous Zinquin staining patterns of PI-positive events.

To assess and capture early changes in [Zn2+]i in response to VP-16, we used two methods of analysis. The first approach evaluated (Fig. 6, A and B) the "balance sheet" for the movement of cells among three nominal fractions: PI-positive, PI-negative/low-[Zn2+]i, and PI-negative/high-[Zn2+]i cells. Discrimination for high vs. low [Zn2+]i was set at the median channel for a Zinquin E-treated control, to avoid prejudice in region setting, with shifts shown relative to the control set at zero. The results show that a shift to the high-[Zn2+]i fraction was always detected for both cell lines at the lowest VP-16 dose (0.25 µM × 3 h) and was significant at 2.5 µM VP-16 (P < 0.05, t-test) for conditions under which apoptosis was not detectable. This increase in PI-negative/high-[Zn2+]i cells was reproducible but was always lost as the VP-16 dose was increased in DoHH2 cells, but not in SU-DHL-4 cells. This loss was due to a movement of cells into the low-[Zn2+]i fraction rather than loss to the nonviable fraction (PI positive). Hence, the analysis presented in Fig. 6, A and B, reflects a demonstrable "switch" from low to high free [Zn2+]i during DNA damage and a subsequent loss of cells from the high fraction reflecting low [Zn2+]i. This approach inherently underestimates event frequency because it is not cumulative with time.


View larger version (24K):
[in this window]
[in a new window]
 
Fig. 6.   Effects of short-term VP-16 exposure (3 h) on the relative change in frequency of DoHH2 (A) and SU-DHL-4 (B) cell subpopulations expressing different [Zn2+]i levels. Intact (PI-negative) cell fractions were defined as having either low [Zn2+]i (i.e., fluorescence intensity <=  median channel for a Zinquin E-treated control) or high [Zn2+]i (i.e., fluorescence intensity > median channel for a Zinquin E-treated control).open circle , PI-negative, low-[Zn2+]i cells; triangle , PI-negative, high-[Zn2+]i cells; , total PI-positive cells. Data are means ± SD for 3 experiments.

The second approach involved the analysis of whole population shifts in [Zn2+]i by using Kolmogorov-Smirnov statistics, measuring the significance of the maximum vertical displacement between two cumulative frequency distributions (49). Figure 7, A-H, demonstrates the heterogeneity in basal levels of free [Zn2+]i and provides evidence of the significance and extent of changes with respect to control distributions. These changes range from an increase in all cells (Fig. 7, E-H), a whole population collapse (Fig. 7, C and D), and a heterogeneous response (Fig. 7, A and B). Figure 7, E-H, shows typical whole population shifts for SU-DHL-4 cells, reporting an increased level of free [Zn2+]i at all doses of VP-16. DoHH2 cells (Fig. 7, A-D) clearly show a significant (P <=  0.001) increase in [Zn2+]i for the low VP-16 dose range. However, at drug doses >2.5 µM, there was a loss of this shift in DoHH2 cells with a fall in [Zn2+]i in the whole population, most noticeably in the high-[Zn2+]i fractions. Overall, there was an early but limited increase in [Zn2+]i detectable in single cells undergoing DNA damage stress, provided apoptosis was not engaged in a permissive cell through the imposition of suprathreshold levels of damage.


View larger version (33K):
[in this window]
[in a new window]
 
Fig. 7.   Kolmogorov-Smirnov (K-S) statistical analyses of Zinquin E-treated DoHH2 (A-D) and SU-DHL-4 cells (E-H) and the changes in [Zn2+]i resulting from a 3-h exposure to different concentrations of VP-16. All K-S analyses showed significant differences in drug-treated cells (P <=  0.001) with D values of 0.15, 0.12, 0.27, 0.51, 0.11, 0.17, 0.13, and 0.18 for 2 sample analyses on data in A-H, respectively. The data show a significant increase in [Zn2+]i at the 0.25-2.5 µM VP-16 dose in DoHH2 cells and at all doses in SU-DHL-4 cells. At doses >2.5 µM VP-16, DoHH2 cells show large reductions in [Zn2+]i.

Extensive analysis of multiple flow cytometry profiles (data not shown) suggested that Zn2+ mobilization, due to the addition of the DNA-damaging agent, is asynchronous with each cell responding at different times and to a different extent. Thus some cells could be increasing free [Zn2+]i while others will have already experienced a collapsed transient, giving rise to mixed populations (Fig. 7, A and B). Furthermore, the inherent heterogeneity in basal free [Zn2+]i means that the true amplitude of the rise and the extent of the collapse for a given cell is unknown.

p53-Independent Induction of Apoptosis by TPEN

The results suggested that early apoptotic engagement may be accompanied by or require a fall in [Zn2+]i and that Zn2+ chelation by TPEN may be capable of providing an apoptotic trigger in DoHH2 cells (Fig. 1B). To test whether apoptosis could be induced by TPEN in p53 mutant cells, apoptotic/cell death changes were quantified by the analysis of nuclear changes detected by laser light scatter (40). Figure 8, A and B, shows the results of a prolonged (18 h) TPEN exposure to permit the development of apoptotic bodies. The data are presented as 1) normal scatter events (i.e., G1 to G2 content DNA with 90° light scatter values within the normal distribution), 2) abnormal scatter events (i.e., G1 to G2 content DNA with 90° light scatter values greater than the normal distribution), and 3) fragmentation events (i.e., >= 0.5× G1 to <1× G1 DNA content). Figure 8A shows that >80% DoHH2 cells show apoptotic characteristics after exposure to >5 µM TPEN. Characteristically, the majority of these cells retained their nuclear DNA content and did not fully fragment even at 250 µM TPEN. Figure 8B shows that SU-DHL-4 cells had a more resistant response with >40% not showing nuclear changes even at the highest dose. However, at >= 5 µM TPEN, ~20% of the SU-DHL-4 cells underwent nuclear changes consistent with apoptosis and a threefold greater level of fragmentation events compared with DoHH2 cells. The data indicate that TPEN exposure (5-50 µM) can induce apoptotic changes in both cell lines. To confirm these observations, we used a complementary method of apoptosis detection, namely, annexin V binding/PI exclusion. The commitment to TPEN-induced apoptosis was found to be rapid and showed similar kinetics for the two cell lines (Fig. 8, C and D). Commitment was followed, after a delay of ~12 h, by essentially the same rate of progression to a nonviable state.


View larger version (23K):
[in this window]
[in a new window]
 
Fig. 8.   Induction of cell death by TPEN in DoHH2 and SU-DHL-4 cell populations analyzed by light scatter changes (A and B) and annexin V binding/PI-exclusion properties (C and D). A and B: triangle , normal scatter events (i.e., G1 to G2 content DNA with 90° light scatter values within the normal distribution); , abnormal scatter events (i.e., G1 to G2 content DNA with 90° light scatter values greater than the normal distribution); open circle , fragmentation events (i.e., >= 0.5 G1 to <G1 DNA content). C and D: time-dependent changes in the frequency of apoptotic (triangle ) and nonviable (black-triangle) cells as defined in Fig. 1 legend. Dashed and solid horizontal lines indicate the range of control values for apoptotic and nonviable cells, respectively.

Enhancement of DNA Damage-Induced Apoptosis by Zn2+ Chelation

The data are consistent with an induction of apoptosis through a fall in intracellular Zn2+ availability and indicate that functional pathways for DNA damage-induced apoptosis are not required. The question arises as to whether Zn2+ chelation-facilitated nuclear fragmentation and DNA damage-induced signals for cell death can interact. If so, cells that are permissive for DNA damage-induced apoptosis should be triggered into apoptosis more readily if there is an imposed reduction in free [Zn2+]i.

This question was tested by using TPEN and the probe Zinquin E itself, because the Zinquin E-cleaved molecules form intracellular complexes with Zn2+ and, hence, act to buffer Zn2+ ions. Figure 9, A and B, shows results for a selected short exposure to VP-16 that did not generate apoptosis or the loss of membrane integrity in either cell type. Zinquin E did not enhance apoptosis in VP-16-treated cells, indicating that the free [Zn2+]i analyses using this probe are not complicated by any induction of cell death pathways by the probe itself during the even shorter period of analysis. The SU-DHL-4 cell line shows a low but significant (<= 7%; P < 0.05) increase in annexin V+ cells following TPEN treatment but no exacerbation in the presence of DNA damage. Because TPEN would abrogate any damage-induced transient in SU-DHL-4 cells, it appears that the transient is not an apoptosis-sparing pathway in DNA-damaged cells. Data for DoHH2 cells confirm that no apoptosis was detectable during a 4-h VP-16 exposure, whereas 22% of TPEN-treated cultures entered apoptosis. We have consistently observed an increased (approximately >1.6-fold for background-corrected samples) entry of DoHH2 cells into apoptosis during this short time period by coincubation with VP-16 (36% cells engaged). Because the data are derived from single cell analyses, we conclude that a reduction of free [Zn2+]i by TPEN provides an enhanced triggering of apoptosis in cells with subthreshold levels of DNA damage and functional genomic stress signaling.


View larger version (21K):
[in this window]
[in a new window]
 
Fig. 9.   Differential induction of apoptotic changes in DoHH2 (A) and SU-DHL-4 (B) cell populations upon 4-h exposure to TPEN, Zinquin E, VP-16, and their combinations. Data are means ± SD for 3 experiments. Shaded bars represent apoptotic cells (lower right quadrant; annexin V+/PI-); open bars represent nonviable cells with disrupted plasma membranes (upper right quadrant; annexin V+/PI+).


    DISCUSSION
TOP
ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
REFERENCES

Here we have shown that early (<= 3 h) increases occur in [Zn2+]i during exposure to biologically relevant VP-16 doses capable of causing an early elevation in p53 levels and induction of p21WAF1 in p53 wild-type cells. This limited early [Zn2+]i transient is also observed in p53 mutant cells, suggesting that intracellular Zn2+ mobilization is p53 independent and does not appear to drive the progression of cells into cycle arrest due to the lack of a dose-response relationship. The early [Zn2+]i transient appears to be more closely related to the immediate effects of DNA damage induction and provides the cell with an environment of increased free intracellular Zn2+ availability during a period of demonstrable p53 stabilization in p53wt cells. However, the transient appears to precede large shifts in accumulation of the downstream effector molecule p21WAF1. This DNA damage-induced free [Zn2+]i transient is also paralleled by, but not dependent on, an upward shift in the apoptosis-promoting bax-to-bcl-2 ratio (23), occurring before the detection of apoptotic changes monitored by annexin V binding. The progressive recruitment of cells into late cell cycle arrest is temporally associated with rapid nuclear fragmentation and occurs preferentially in those cells showing elevated levels of p53 protein. Our results show that exposure of DoHH2 cells to higher VP-16 dose levels, eventually leading to arrest and DNA fragmentation, induces an early fall in free [Zn2+]i. The collapse of the free [Zn2+]i transient is expressed in an increasing number of cells as the VP-16 dose, and therefore the probability of recruitment into apoptosis, increases.

Enforcing a fall in free [Zn2+]i by using the chelator TPEN in the p53 wild-type cell line, permissive for the engagement of apoptosis in the presence of DNA damage signals, caused a rapid commitment to apoptosis. This suggests that a reduction in free [Zn2+]i can act as an apoptotic trigger as observed by others (9, 29, 33, 35, 44). Our results also indicate that VP-16 cotreatment of p53 wild-type cells can exacerbate this TPEN-induced early commitment to apoptosis. This suggests that low free [Zn2+]i can permit normally subthreshold levels of VP-16-induced DNA damage to trigger apoptosis. The p53 mutant cell line shows a highly muted apoptotic response to VP-16-induced DNA damage. However, the difference in the TPEN apoptotic response between the two cell lines is far less pronounced. TPEN exposure results in up to 40% of SU-DHL-4 cells progressing to limited nuclear fragmentation, with kinetics very similar to those for DoHH2 cells. Thus it appears that following a short or sustained reduction in free [Zn2+]i, the p53 mutant cell line can also engage elements of the apoptotic pathway, reenforcing the concept that low free [Zn2+]i per se can trigger DNA fragmentation. The early free [Zn2+]i transient in VP-16-treated SU-DHL-4 cells does not collapse as the drug concentration increases, suggesting that high levels of DNA damage per se, as evidenced by the increasing expression of G2 arrest, do not cause a fall in free [Zn2+]i. The data are consistent with the collapse being a feature of cells destined to undergo DNA fragmentation within the apoptotic response.

In this model, [Zn2+]i homeostasis is an integral part of the DNA fragmentation response. Accordingly, enforcing increased levels of free intracellular Zn2+ might be expected to inhibit cell death (10). The presence of Zn2+ in culture medium can interfere with DNA damage-associated apoptosis in UVB-irradiated HaCaT keratinocytes (2) and cisplatin-treated HeLa cells (27). On the other hand, concentrations of extracellular Zn2+ (500-1,000 µM) have been used to block, at least short-term, glucocorticoid-induced apoptosis in thymocytes, whereas lower or more physiological concentrations of Zn2+ (80-200 µM) appear to be able to induce cell death (18). The balance between the basal level of free intracellular Zn2+ in a cell, the extent of stress-induced release of Zn2+, and the early Zn2+-consumption requirements of a cell for the pursuit of apoptosis would potentially affect the efficiency with which apoptosis is inhibited by extracellular Zn2+. The setting of the balance for free [Zn2+]i would also contribute to the cell cycle heterogeneity, intercell line variation, and the complex effects of chelating agents and extracellular Zn2+.

It appears that cells may integrate Zn2+ homeostasis with apoptosis at the level of caspase function, providing control over internucleosomal fragmentation and the cleavage of the DNase inhibitor ICAD. Zn2+ depletion-induced apoptosis by TPEN is associated with activation of caspases-3, -8, and -9 and cleavage of target proteins (11). Additionally, Zn2+ addition partially inhibits caspase-3 activation, but not caspase-8 and -9 cleavage, in VP-16-treated HeLa cells (11), suggesting that the impact of upstream DNA damage-originating apoptotic signals may be affected by intracellular Zn2+ availability at the level of caspase-3. In cells undergoing oxidative stress, limited caspase-3 activation can occur with a resumption of activation and progression to internucleosomal fragmentation upon mild Zn2+ chelation (26). Such observations suggest that in the presence of strong or persistent apoptotic signals, relatively small-scale changes in free [Zn2+]i may play a critical role in the full engagement and active pursuance of apoptosis. Accordingly the induction of apoptosis by TPEN in both DoHH2 and SU-DHL-4 cells could result from activation of caspase-3 and serves to demonstrate the functional integrity of the downstream apoptotic cascade even in the p53 mutant cell line. We suggest that the observed collapse of the [Zn2+]i transient in high-dose VP-16-treated DoHH2 cells results in a partial activation of caspase-3, permitting the rapid and extensive triggering of apoptosis through DNA damage signaling. The early enhancement of VP-16-induced apoptosis by TPEN in the p53 wild-type cell line could arise from the reenforcement of upstream proapoptotic signals acting at a partially activated caspase-3.

The initial increase in [Zn2+]i would provide resistance to apoptotic signals through caspase-3 partial inactivation while the cell attempts to initiate checkpoint activation. Our VP-16 cell cycle study indicates that both lymphoma cell lines show similar potential for cycle arrest, except that events in DoHH2 cells are truncated by the engagement of apoptosis. Other studies have shown that TPEN treatment of epithelial cells results in the early (1-2 h) activation of caspase-3 and in the rapid cleavage of p21WAF1/CIP1 to a 15-kDa fragment before further degradation and the appearance of morphologic changes characteristic of apoptosis (9). During the p53-mediated response to gamma radiation-induced DNA damage, p21WAF1/CIP1 is rapidly induced but selectively cleaved, probably by a caspase-3-like activity (19). Cleavage abrogates interaction with proliferating cell nuclear antigen (PCNA) potentially interfering with normal PCNA-dependent repair and checkpoint function. A Zn2+ transient may provide an apoptosis-inhibiting period during which the cleavage of target cell cycle inhibitor proteins is blocked, permitting the progressive recruitment of cells into cell cycle arrest.

Zn2+ transients have been observed in cells undergoing differentiation in culture (35), and previous work using Zinquin has suggested that apoptosis, arising spontaneously or following induction by DNA-damaging agents, can be accompanied by a release of Zn2+ from intracellular stores or MTs (50). In assessing the reactive changes in shifts in free [Zn2+]i induced by VP-16, it is clear that they are highly drug dose dependent. This reflects the sharp transition between the effects of low doses in permitting progression to G2/M arrest and the effects of higher doses (>0.5 µM) in trapping cells in S phase and inducing an earlier commitment to apoptosis (41) as threshold levels of damage are surpassed (32). The ability to observe free [Zn2+]i transients is severely restricted by the rapidity with which permissive cells commit to apoptosis and thereupon reduce available intracellular Zn2+ levels. We have observed that cells with compromised membrane integrity also show elevated Zinquin fluorescence. This potential artifact was specifically excluded from our measurements.

The origin of the increases in free [Zn2+]i is assumed to be preexisting intracellular MT pools (4). The fall in free [Zn2+]i could represent transport, sequestration, or use by free Zn2+-requiring pathways. However, reactive changes in free [Zn2+]i may be compartmentalized. Punctate staining of cells with Zinquin has been noted before (28) and was demonstrated here for both control and extracellular Zn2+-loaded lymphoma cells by using the novel approach of two-photon excitation of the Zn2+ probe. It has been suggested that the intravesicular pools of Zn2+ ("zincosomes"; Ref. 10) are associated with cytoskeletal actin and protein kinase C molecules. With the use of Zn2+-loaded calibration samples, the mean free [Zn2+]i for control populations was estimated to be ~100 pmol/106 SU-DHL-4 cells with nearly a fivefold difference between the lowest and highest [Zn2+]i levels. This compares with a previous estimate of the average content of labile Zn2+ in human leukemic lymphocytes of ~20 pmol/106 cells (50).

We conclude that the early phases of stress-induced p53-dependent gene activation occur within a readily saturated environment of elevated free [Zn2+]i and that breaching thresholds of DNA damage and engaging apoptosis in permissive cells are preceded by a necessary reduction of free [Zn2+]i. The responses of the p53 mutant cell line suggest that DNA damage or drug presence per se is not responsible for the reduction in free [Zn2+]i; rather, it is likely to arise from the responses to suprathreshold levels of genomic stress. The loss of [Zn2+]i may reflect the binding of the cation by metal-binding protein(s) required for nuclear destruction. Collapse of the transient may release destruction effector molecules such as caspases, normally inhibited by Zn2+-requiring IAPs (32, 48), supported by the transient increase in Zn2+ following subapoptotic levels of stress. The study reveals that determination of true causality in the linkage between free [Zn2+]i changes and cell death/cycle delay processes is problematic. Whether or not drug-reactive shifts in labile Zn2+ actually affect the activity of pathways that use Zn2+-requiring proteins depends on pool location, protein-metal cation equilibria, the time frame within which such shifts occur, and the functional integrity of the zinc proteins and their downstream effectors. The development of compartment-specific ratiometric probes for [Zn2+]i and molecular structures for the caged release of Zn2+ would greatly aid such investigations.

Our findings have implications for understanding the effects of Zn2+-sequestering molecules such as MTs and exogenously supplied chelating agents on the two main competing responses of human cells to genomic stress that can affect population dynamics, namely, the resolution of cell cycle arrest vs. active progression to cell death. We suggest that heterogeneity in intracellular Zn2+ levels within normal tissues and, indeed, tumor cell populations could act to limit their ability to collapse free intracellular Zn2+ concentrations and efficiently pursue cell death responses to physiologically derived signals or those arising from imposed genomic stress.

The dynamic processes that lead to Zn2+ transients and collapse are not clear, although the current study emphasizes the event heterogeneity and rapidity. If such events occur in parallel with the initiation of cycle arrest and the transduction of apoptotic signals, then their impact on the modulation of caspase activity or p53-driven processes will depend on when they occur. Studies involving enforced changes of Zn2+ levels by chelation cannot reproduce or mimic the temporal linking of such Zn2+ changes with those signals. Future studies should address the nature of Zn2+ profiles in single cells containing additional reporters to provide the temporal linking to damage signaling events.

The concept that apoptosis is engaged as thresholds of DNA damage are surpassed has been discussed previously and describes an important strategy in mammals for coping with injury and tissue organization (32). The implication of the current study is that free [Zn2+]i transients imposed by physiological changes could determine the thresholds for engaging cell death pathways under biologically relevant levels of persistent genomic stress.


    ACKNOWLEDGEMENTS

This work was supported by grants from the Medical Research Council (UK) and the Joint Research Equipment Initiative (UK).


    FOOTNOTES

Address for reprint requests and other correspondence: P. J. Smith, Dept. of Pathology, Univ. of Wales College of Medicine, Heath Park, Cardiff CF14 4XN, UK (E-mail: smithpj2{at}cf.ac.uk).

The costs of publication of this article were defrayed in part by the payment of page charges. The article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.

March 6;10.1152/ajpcell.00439.2001

Received 13 September 2001; accepted in final form 27 February 2002.


    REFERENCES
TOP
ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
REFERENCES

1.   Abdel-Mageed, AB, and Agrawal KC. Antisense down-regulation of metallothionein induces growth arrest and apoptosis in human breast carcinoma cells. Cancer Gene Ther 4: 199-207, 1997[ISI][Medline].

2.   Ahn, YH, Kim YH, Hong SH, and Koh JY. Depletion of intracellular zinc induces protein synthesis-dependent neuronal apoptosis in mouse cortical culture. Exp Neurol 154: 47-56, 1998[ISI][Medline].

3.   Anstey, A, Marks R, Long C, Navabi H, Pearse A, Wynford Thomas D, and Jasani B. In-vivo photoinduction of metallothionein in human skin by ultraviolet-irradiation. J Pathol 178: 84-88, 1996[ISI][Medline].

4.   Berendji, D, Kolb-Bachofen V, Meyer KL, Grapenthin O, Weber H, Wahn V, and Kroncke KD. Nitric oxide mediates intracytoplasmic and intranuclear zinc release. FEBS Lett 405: 37-41, 1997[ISI][Medline].

5.   Berg, JM. Zinc fingers, and other metal-binding domains. Elements for interactions between macromolecules. J Biol Chem 265: 6513-6516, 1990[Free Full Text].

6.   Beyersmann, D, and Hechtenberg S. Cadmium, gene regulation, and cellular signalling in mammalian cells. Toxicol Appl Pharmacol 144: 247-261, 1997[ISI][Medline].

7.   Borresen-Dale, AL, Lothe RA, Meling GI, Hainaut P, Rognum TO, and Skovlund E. TP53 and long-term prognosis in colorectal cancer: mutations in the L3 zinc-binding domain predict poor survival. Clin Cancer Res 4: 203-210, 1998[Abstract].

8.   Bunz, F, Dutriaux A, Lengauer C, Waldman T, Zhou S, Brown JP, Sedivy JM, Kinzler KW, and Vogelstein B. Requirement for p53 and p21 to sustain G2 arrest after DNA damage. Science 282: 1497-1501, 1998[Abstract/Free Full Text].

9.   Chai, F, Truong-Tran AQ, Evdokiou A, Young GP, and Zalewski PD. Intracellular zinc depletion induces caspase activation and p21Waf1/Cip1 cleavage in human epithelial cell lines. J Infect Dis 182 Suppl1: S85-S92, 2000[ISI][Medline].

10.   Chai, F, Truong-Tran AQ, Ho LH, and Zalewski PD. Regulation of caspase activation and apoptosis by cellular zinc fluxes and zinc deprivation: a review. Immunol Cell Biol 77: 272-278, 1999[ISI][Medline].

11.   Chimienti, F, Seve M, Richard S, Mathieu J, and Favier A. Role of cellular zinc in programmed cell death: temporal relationship between zinc depletion, activation of caspases, and cleavage of Sp family transcription factors. Biochem Pharmacol 62: 51-62, 2001[ISI][Medline].

12.   Coleman, JE. Zinc proteins: enzymes, storage proteins, transcription factors, and replication proteins. Annu Rev Biochem 61: 897-946, 1992[ISI][Medline].

13.   Coyle, P, Zalewski PD, Philcox JC, Forbes IJ, Ward AD, Lincoln SF, Mahadevan I, and Rofe AM. Measurement of zinc in hepatocytes by using a fluorescent probe, Zinquin: relationship to metallothionein and intracellular zinc. Biochem J 303: 781-786, 1994[ISI][Medline].

14.   Eid, H, Geczi L, Bodrogi I, Institoris E, and Bak M. Do metallothioneins affect the response to treatment in testis cancers? J Cancer Res Clin Oncol 124: 31-36, 1998[ISI][Medline].

15.   El-Deiry, WS, Harper JW, O'Connor PM, Velculescu VE, Canman CE, Jackman J, Pietenpol JA, Burrell M, Hill DE, Wang Y, Wiman KG, Mercer WE, Kastan MB, Kohn KW, Elledge SJ, Kinzler KW, and Vogelstein B. WAF1/CIP1 is induced in p53-induced G1 arrest and apoptosis. Cancer Res 54: 1169-1174, 1994[Abstract].

16.   Epstein, AL, Levy R, Kim H, Henle W, Henle G, and Kaplan HS. Biology of the human malignant lymphomas. IV. Functional characterization of ten diffuse histiocytic lymphoma cell lines. Cancer 42: 2379-2391, 1978[ISI][Medline].

17.   Fanzo, JC, Reaves SK, Cui L, Zhu L, Wu JYJ, Wang YR, and Lei KY. Zinc status affects p53, gadd45, and c-fos expression and caspase-3 activity in human bronchial epithelial cells. Am J Physiol Cell Physiol 281: C751-C757, 2001[Abstract/Free Full Text].

18.   Fraker, PJ, and Telford WG. A reappraisal of the role of zinc in life and death decisions of cells. Proc Soc Exp Biol Med 215: 229-236, 1997[Abstract].

19.   Gervais, JL, Seth P, and Zhang H. Cleavage of CDK inhibitor p21Cip1/Waf1 by caspases is an early event during DNA damage-induced apoptosis. J Biol Chem 273: 19207-19212, 1998[Abstract/Free Full Text].

20.   Hirayama, Y. Histochemical localization of zinc and copper in rat ocular tissues. Acta Histochem 89: 107-111, 1990[ISI][Medline].

21.   Kluin-Nelemans, HC, Limpens J, Meerabux J, Beverstock GC, Jansen JH, de Jong D, and Kluin PM. A new non-Hodgkin's B-cell line (DoHH2) with a chromosomal translocation t(14,18)(q32,q21). Leukemia 5: 221-224, 1991[ISI][Medline].

22.   Kondo, Y, Rusnak JM, Hoyt DG, Settineri CE, Pitt BR, and Lazo JS. Enhanced apoptosis in metallothionein null cells. Mol Pharmacol 52: 195-201, 1997[Abstract/Free Full Text].

23.   Korsmeyer, SJ. BCL-2 gene family and the regulation of programmed cell death. Cancer Res 59: 1693-1700, 1999.

24.   Laemmli, UK. Cleavage of structural proteins during the assembly of the head of bacteriophage T4. Nature 227: 680-685, 1970[ISI][Medline].

25.   Magneson, GR, Puvathingal JM, and Ray WJ. The concentrations of free Mg2+ and free Zn2+ in equine blood plasma. J Biol Chem 262: 11140-11148, 1987[Abstract/Free Full Text].

26.   Marini, M, Frabetti F, Canaider S, Dini L, Falcieri E, and Poirier GG. Modulation of caspase-3 activity by zinc ions and by the cell redox state. Exp Cell Res 266: 323-332, 2001[ISI][Medline].

27.   Marini, M, and Musiani D. Micromolar zinc affects endonucleolytic activity in hydrogen peroxide-mediated apoptosis. Exp Cell Res 239: 393-398, 1998[ISI][Medline].

28.   Palmiter, RD, Cole TB, and Findley SD. ZnT-2, a mammalian protein that confers resistance to zinc by facilitating vesicular sequestration. EMBO J 15: 1784-1791, 1996[Abstract].

29.   Parat, MO, Richard MJ, Pollet S, Hadjur C, Favier A, and Beani JC. Zinc and DNA fragmentation in keratinocyte apoptosis: its inhibitory effect in UVB irradiated cells. J Photochem Photobiol B 37: 101-106, 1997[ISI][Medline].

30.   Peck, EJ, and Ray WJ. Metal complexes of phosphoglucomutase in vivo. J Biol Chem 246: 1160-1167, 1971[Abstract/Free Full Text].

31.   Perry, ME, Piette J, Zawadzki JA, Harvey D, and Levine AJ. The mdm-2 gene is induced in response to uv-light in a p53-dependent manner. Proc Natl Acad Sci USA 90: 11623-11627, 1993[Abstract].

32.   Rich Allen, T, RL, and Wyllie AH. Defying death after DNA damage. Nature 407: 777-783, 2000[ISI][Medline].

33.   Sakabe, I, Paul S, Dansithong W, and Shinozawa T. Induction of apoptosis in Neuro-2A cells by Zn2+ chelating. Cell Struct Funct 23: 95-99, 1998[ISI][Medline].

34.   Sambrook, F, Fritsch EF, and Maniatis T. Detection and analysis of proteins expressed from cloned genes. In: Molecular Cloning: A Laboratory Manual. Cold Spring Harbor, New York: Cold Spring Harbor Laboratories, 1989.

35.   Schmidt, C, and Beyersmann D. Transient peaks in zinc and metallothionein levels during differentiation of 3T.3L1 cells. Arch Biochem Biophys 364: 91-98, 1999[ISI][Medline].

36.   Simons, TJB Intracellular free zinc and zinc buffering in human red blood cells. J Membr Biol 123: 63-71, 1991[ISI][Medline].

37.   Smith, PJ, Blunt N, Wiltshire M, Hoy T, Teesdale-Spittle P, Craven MR, Watson JV, Amos WB, Errington RJ, and Patterson LH. Characteristics of a novel deep red/infra red fluorescent cell permeant DNA probe, DRAQ5, in intact human cells analysed by flow cytometry, confocal and multiphoton microscopy. Cytometry 40: 280-291, 2000[ISI][Medline].

38.   Smith, PJ, and Jones CJ. p53 and the integrated cellular response to DNA damage. In: DNA Recombination and Repair. Oxford, UK: IRL, 1999, p. 202-231.

39.   Smith, PJ, Soues S, Gottlieb T, Falk SJ, Watson JV, Osborne RJ, and Bleehen NM. Etoposide-induced cell cycle delay and arrest-dependent modulation of DNA topoisomerase II in small cell lung cancer cells. Br J Cancer 70: 914-921, 1994[ISI][Medline].

40.   Smith, PJ, Wiltshire M, Chin SF, Rabbitts P, and Souès S. Cell cycle checkpoint evasion and protracted cell cycle arrest in X-irradiated small cell lung carcinoma cells. Int J Radiat Biol 75: 1137-1147, 1999[ISI][Medline].

41.   Souès, S, Wiltshire M, and Smith PJ. Differential sensitivity to etoposide (VP-16)-induced S phase delay in a panel of small cell lung carcinoma cell lines with G1/S-phase checkpoint dysfunction. Cancer Chemother Pharmacol 47: 133-140, 2001[ISI][Medline].

42.   Towbin, H, Staehelin T, and Gordon J. Electrophoretic transfer of proteins from polyacrylamide gels to nitrocellulose sheets: Procedure and some applications. Proc Natl Acad Sci USA 76: 4350-4354, 1979[Abstract].

43.   Vallee, BL, and Auld DS. Zinc: biological functions and coordination motifs. Accts Chem Res 26: 543-551, 1993.

44.   Velazquez, M, Maldonado V, and Melendez-Zajgla J. Cisplatin-induced apoptosis of HeLa cells. Effect of RNA and protein synthesis inhibitors, Ca2+ chelators and zinc. J Exp Clin Cancer Res 17: 277-284, 1998[ISI][Medline].

45.   Verhaegh, GW, Parat MO, Richard MJ, and Hainaut P. Modulation of p53 protein conformation and DNA-binding activity by intracellular chelation of zinc. Mol Carcinog 21: 205-214, 1998[ISI][Medline].

46.   Vermes, I, Haanen C, Steffens-Nakken H, and Reutelingsperger CA. novel assay for apoptosis: flow cytometric detection of phosphatidylserine expression on early apoptotic cells using fluorescein-labeled annexin V. J Immunol Methods 184: 39-51, 1995[ISI][Medline].

47.   Winters, ZE, Ongkeko WM, Harris AL, and Norbury CJ. P53 regulates Cdc2 independently of inhibitory phosphorylation to reinforce radiation-induced G2 arrest in human cells. Oncogene 17: 673-684, 1998[ISI][Medline].

48.   Wu, G, Chai J, Suber TL, Wu JW, Du C, Wang X, and Shi Y. Structural basis of I.A.P recognition by Smac/DIABLO. Nature 408: 1008-1012, 2000[ISI][Medline].

49.   Young, IT. Proof without prejudice: use of the Kolmogorov-Smirnov test for the analysis of histograms from flow systems and other sources. J Histochem Cytochem 25: 935-941, 1977[Abstract].

50.   Zalewski, PD, Forbes IJ, and Betts WH. Correlation of apoptosis with change in intracellular labile Zn(II) using Zinquin [(2-methyl-8-p-toluenesulphonamido-6-quinolyloxy)acetic acid], a new specific fluorescent probe for Zn(II). Biochem J 296: 403-408, 1993[ISI][Medline].

51.   Zalewski, PD, Forbes IJ, Seamark RF, Borlinghaus R, Betts WH, Lincoln SF, and Ward AD. Flux of intracellular labile zinc during apoptosis (gene-directed cell death) revealed by a specific chemical probe, Zinquin. Chem Biol 1: 153-161, 1994[Medline].


Am J Physiol Cell Physiol 283(2):C609-C622
0363-6143/02 $5.00 Copyright © 2002 the American Physiological Society




This Article
Abstract
Full Text (PDF)
All Versions of this Article:
283/2/C609    most recent
00439.2001v1
Alert me when this article is cited
Alert me if a correction is posted
Citation Map
Services
Email this article to a friend
Similar articles in this journal
Similar articles in ISI Web of Science
Similar articles in PubMed
Alert me to new issues of the journal
Download to citation manager
Search for citing articles in:
ISI Web of Science (6)
Google Scholar
Articles by Smith, P. J.
Articles by Errington, R. J.
Articles citing this Article
PubMed
PubMed Citation
Articles by Smith, P. J.
Articles by Errington, R. J.


HOME HELP FEEDBACK SUBSCRIPTIONS ARCHIVE SEARCH TABLE OF CONTENTS
Visit Other APS Journals Online