The giant cytosolic protease tripeptidyl peptidase II (TPPII) was recently proposed to play a role in the DNA damage response. Shown were nuclear translocation of TPPII after γ-irradiation, lack of radiation-induced p53 stabilization in TPPII-siRNA–treated cells, and complete tumor regression in mice after γ-irradiation when combined with TPPII-siRNA silencing or a protease inhibitor reported to inhibit TPPII. This suggested that TPPII could be a novel target for tumor radiosensitization and prompted us to study radiation responses using TPPII-knockout mice. Neither the sensitivity to total body irradiation nor the radiosensitivity of resting lymphoid cells, which both strongly depend on p53, was altered in the absence of TPPII. Functional integrity of p53 in TPPII-knockout cells is further shown by a proper G1 arrest and by the accumulation of p53 and its transcriptional targets, p21, Bax, and Fas, on γ-irradiation. Furthermore, we could not confirm radiation-induced nuclear translocation of TPPII. Nevertheless, after γ-irradiation, we found slightly increased mitotic catastrophe of TPPII-deficient primary fibroblasts and increased apoptosis of TPPII-deficient activated CD8+ T cells. The latter was accompanied by delayed resolution of the DNA double-strand break marker γH2AX. This could, however, be due to increased apoptotic DNA damage rather than reduced DNA damage repair. Our data do not confirm a role for TPPII in the DNA damage response based on nuclear TPPII translocation and p53 stabilization but nevertheless do show increased radiation-induced cell death of selected nontransformed cell types in the absence of the TPPII protease. [Cancer Res 2009;69(8):3325–31]

With a molecular mass of 6 MDa, tripeptidyl peptidase II (TPPII) forms the largest protease complex in eukaryotic cells (1). TPPII was discovered as a cytosolic aminotripeptidase that rapidly degraded synthetic oligopeptides (2). Meanwhile, TPPII complexes have also been shown to cleave short polypeptides by exopeptidase as well as trypsin-like endopeptidase activity (35). Yet, whether TPPII contributes to the degradation of full-length proteins or has nonproteolytic functions has remained elusive. A major cellular function of TPPII is probably in cytosolic proteolysis downstream of proteasomes (2).

In several reports, TPPII has been linked with proliferation and survival of lymphoma cells, particularly under conditions of cellular stress, such as continuous proteasome inhibition (3, 6), overexpression of c-Myc (7), lack of nutrients, or high in vivo tumor cell proliferation (8). There is also evidence for a role of TPPII in proliferation and survival of nonlymphoid tumor cells (9, 10). A role in the regulation of centrosome homeostasis and mitotic fidelity has also been proposed (9, 10). These studies were conducted with protease inhibitors, TPPII-specific siRNA, or overexpression of TPPII.

Our initial analysis of TPPII-knockout (KO) mice revealed a role of TPPII for proliferation and survival of several primary cell types (11). TPPII-deficient primary fibroblasts and T-cell receptor–stimulated CD8+ T cells exhibit increased premature cellular senescence, and proliferating CD8+ T cells, in addition, have increased apoptosis. Increased apoptosis was also found for the small population of rapidly proliferating double-negative thymocytes. In vivo, increased cell death in these cell types contributes to premature immunosenescence and perhaps also to the observed premature death of elderly TPPII-KO mice. Increased apoptosis was found to be associated with alterations in the activation of NF-κB.

Recently, a role of TPPII in the cellular response to γ-irradiation and in the in vivo radiation response of tumors was reported by Hong and colleagues (12). Major findings were (a) up-regulation and translocation of TPPII to the nucleus on γ-irradiation; (b) failure of TPPII-siRNA-expressing lymphoma cells to stabilize p53 and consequently to halt the cell cycle in the G1 phase on γ-irradiation; and (c) complete remission of subcutaneous murine EL4 lymphomas on low-dose total body irradiation combined with either intratumoral expression of TPPII-specific siRNA or systemic administration of a subtilase inhibitor, which was reported by Hong and colleagues (12) to inhibit TPPII. These findings suggested not only an unexpected role of TPPII in the DNA damage response but also a potential suitability of TPPII inhibitors as radiosensitizers for clinical use in the radiotherapy of tumors. Hence, we deemed it important to explore the potential role of TPPII in cellular and systemic radiation responses with the help of our TPPII-KO mice.

Mice. Generation of TPPII-KO mice was described previously (11). p53-deficient mice (C57/BL6, The Jackson Laboratory; ref. 13) were obtained from the Max-Planck Institute of Immunobiology in Freiburg. All animal experiments were done in compliance with the guidelines of the University of Freiburg.

Antibodies. An antiserum against the NH2-terminal region of TPPII was produced by peptide immunization in rabbits, and the crude antiserum was affinity purified with the antigenic peptide coupled to a Sepharose column. Another anti-TPPII antibody, which is directed against an internal region of TPPII (E-17), was purchased from Santa Cruz. Anti–phospho-ATM, anti-Bax, anti-p21, and anti-actin antibodies were also from Santa Cruz. Anti–phospho-p53 (Ser15) and anti–phospho-γH2AX (Ser139) were from Cell Signaling, anti-Fas was from BioVision, and anti-Sp1 from BD Bioscience. Anti-p53 was purchased from Santa Cruz or Cell Signaling. Secondary anti–horseradish peroxidase antibodies were all from Dianova. Secondary antibody for immunofluorescence experiments was anti–rabbit Alexa Fluor 546 from Invitrogen.

Cell culture. In all experiments, DMEM supplemented with 10% FCS, l-glutamine (2 mmol/L), 100 μmol/L 2-mercaptoethanol, 10 mL/L NEM (×100), and penicillin/streptomycin (100 units/mL) was used. Cells were incubated at 37°C and 8.5% CO2.

Generation of primary mouse skin fibroblasts. Primary fibroblasts were generated from the skin of newborn mice. The skin was cut into small pieces and trypsinized for 1 h at 37°C and 8.5% CO2. Then digested skin pieces were collected, washed with medium, and cultured as described above.

Activation of CD8+ T cells. Splenic and lymph node CD8+ T cells were isolated with a CD8+ T-cell isolation kit (Miltenyi) and then incubated in medium supplemented with phorbol 12-myristate 13-acetate (PMA; 10 ng/mL) and ionomycin (500 ng/mL), both from Sigma, or anti-CD3 (5 μg/mL) plus anti-CD28 (2 μg/mL), both from eBioscience.

Total body irradiation. Age-matched mice were irradiated with 7 Gy delivered from a Gammacell 40 137Cs laboratory irradiator. Survival was monitored for 30 d.

Apoptosis assays. Nonactivated splenic and thymic lymphocytes or activated splenic and lymph node CD8+ lymphocytes were suspended in medium (4 × 106/mL for spleen cells and 2 × 106/mL for thymocytes), irradiated, and, at the time points indicated, stained with Annexin V and propidium iodide using an Annexin V-FITC Kit from BD Pharmingen. Apoptosis was measured by flow cytometry on a Cytomics FC 500 instrument from Beckman Coulter. To detect apoptosis in primary fibroblasts, cells were irradiated and apoptosis was measured 48 h later.

Assessment of mitotic catastrophe. Primary fibroblasts at passage 2 were seeded on coverslips. After 24 h, cells were irradiated and, 48 h later, fixed and stained with 4′-6-diamidino-2-phenylindole (DAPI) and analyzed under a BX41 fluorescence microscope from Olympus equipped with a digital camera CC-12 soft imaging system (U-CMAD3, Olympus).

Cell cycle analyses. Primary (passage 2) fibroblasts and isolated CD8+ T cells from spleen and lymph nodes, which had been activated with PMA and ionomycin, were irradiated, fixed at the indicated time points with 70% ethanol, and stored overnight at −20°C. Cells were then washed and incubated with propidium iodide (50 μg/mL) and RNase (100 μg/mL) for 2 h at 4°C. After washing, the cells were analyzed for DNA content by flow cytometry.

Analysis of the G1 checkpoint. Activated spleen and lymph node CD8+ T cells were irradiated, and 8 and 16 h later, 5-bromo-2-deoxyuridine (BrdUrd) was added to a final concentration of 10 μmol/L for 30 min. Cells were then washed with PBS and stained with the FITC BrdU Flow Kit from BD Pharmingen and analyzed for BrdUrd incorporation and 7-amino-actinomycin D (7-AAD) DNA staining by flow cytometry.

Immunoblot analysis. Cell lysates were prepared in radioimmunoprecipitation assay (RIPA) lysis buffer [1% NP40, 1% sodium deoxycholate, 0.1% SDS, 0.15 mol/L NaCl, 0.01 mol/L sodium phosphate (pH 7.2), 2 mmol/L EDTA, 50 mmol/L sodium fluoride, and 1 mmol/L sodium vanadate] supplemented with a protease inhibitor cocktail (Complete from Roche). Then, 50 μg of cell lysate per lane were separated by SDS-PAGE. The blots were probed with the indicated antibodies and developed by enhanced chemiluminescence (Amersham Biosciences).

Immunofluorescence staining. Mouse EL4 cells were irradiated and spun onto slides using a Cytospin centrifuge (Thermo Electron). The cells were then fixed with ice-cold methanol for 5 min at −20°C. Thereafter, the cells were permeabilized with ice-cold acetone for 10 s. After blocking (2% bovine serum albumin and 5% goat serum in PBS for 5 min at room temperature), the cells were incubated with anti-TPPII antibodies for 20 min at room temperature, followed by incubation with Alexa Fluor 546–labeled secondary antibodies for 20 min at room temperature. Nuclei were counterstained with DAPI, and cells analyzed using a BX41 fluorescence microscope (Olympus) equipped with the digital camera CC-12 soft imaging system U-CMAD3 at 100-fold magnification.

Preparation of cytosolic and nuclear fractions. For subcellular fractionation, cells were washed and incubated for 5 min on ice in a hypotonic lysis buffer [25 mmol/L HEPES (pH 7.4), 2 mmol/L EGTA, 2 mmol/L MgCl2, and 2 mmol/L DTT] supplemented with a protease inhibitor cocktail (Complete from Roche). Cells were then lysed by two freezing/thawing cycles in liquid nitrogen. Nuclei were obtained by centrifugation at 500 × g (5 min 4°C), washed several times in hypotonic lysis buffer, and lysed in RIPA buffer (described above). The postnuclear supernatant was spun at 13,000 rpm for 15 min at 4°C, and the supernatant collected as the cytosolic fraction.

Statistical analyses. All data are presented as mean ± SD and analyzed by two-tailed Student's t test with unequal variance. P < 0.05 was considered significant.

Total body radiosensitivity and radiosensitivity of lymphoid cells. The radiosensitivity of immature hematopoietic cells and resting mature lymphoid cells is strongly dependent on p53. Accordingly, p53-KO mice survive doses of total body irradiation up to 9 or 10 Gy, which in wild-type (WT) animals cause the development of lethal bone marrow depletion—hematopoietic syndrome (1417). Because TPPII has been proposed to be important for the γ-radiation–induced stabilization of p53 (12), we first compared the total body radiosensitivity of TPPII WT and KO mice. As seen in Fig. 1A, we did not observe any better survival of TPPII-KO compared with WT mice on sublethal total body irradiation with 7 Gy. Rather, the survival of the KO mice was slightly worse, although the difference was not statistically significant.

Figure 1.

Total body irradiation and sensitivity of resting and activated lymphoid cells to ionizing radiation. A, Kaplan-Meier curve for survival of WT and TPPII-KO mice after total body irradiation (TBI). Six- to eight-week-old WT and KO mice (n = 14 each genotype) were subjected to 7-Gy total body irradiation and survival was recorded for 30 d. B, thymocytes from WT and TPPII-KO or p53 KO mice were irradiated with 5 Gy (left) or with varying doses (right) directly ex vivo. At the indicated time points, apoptosis was measured by flow cytometry after staining with Annexin V and propidium iodide. C, primary TPPII WT and KO splenocytes were irradiated with 5 Gy directly ex vivo. At the indicated time points, apoptosis was measured as described in B. D, isolated spleen and lymph node CD8+ WT and TPPII-KO T cells were activated in medium supplemented with PMA (10 ng/mL) and ionomycin (500 ng/mL). After 48 h, cells were irradiated with 5 Gy and apoptosis was assessed another 20 h later as described in B. Each of the experiments in B to D was done two to four times.

Figure 1.

Total body irradiation and sensitivity of resting and activated lymphoid cells to ionizing radiation. A, Kaplan-Meier curve for survival of WT and TPPII-KO mice after total body irradiation (TBI). Six- to eight-week-old WT and KO mice (n = 14 each genotype) were subjected to 7-Gy total body irradiation and survival was recorded for 30 d. B, thymocytes from WT and TPPII-KO or p53 KO mice were irradiated with 5 Gy (left) or with varying doses (right) directly ex vivo. At the indicated time points, apoptosis was measured by flow cytometry after staining with Annexin V and propidium iodide. C, primary TPPII WT and KO splenocytes were irradiated with 5 Gy directly ex vivo. At the indicated time points, apoptosis was measured as described in B. D, isolated spleen and lymph node CD8+ WT and TPPII-KO T cells were activated in medium supplemented with PMA (10 ng/mL) and ionomycin (500 ng/mL). After 48 h, cells were irradiated with 5 Gy and apoptosis was assessed another 20 h later as described in B. Each of the experiments in B to D was done two to four times.

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Intact p53 function in TPPII-deficient cells is also suggested by the virtually indistinguishable strong apoptosis observed for TPPII WT and KO thymocytes or splenocytes irradiated directly ex vivo (Fig. 1B and C). In contrast, both p53-deficient thymocytes and splenocytes resisted even very high radiation doses, as described in the literature (13, 17). The presence of TPPII in WT lymphoid cells and its absence in the TPPII-KO cells were verified by Western blotting (Supplementary Fig. S1). Most primary thymocytes and lymphocytes are resting under normal conditions. Mitogenic activation greatly inhibits T-cell apoptosis in response to DNA damage (17), and it is therefore interesting that we found a relatively strong increase in apoptosis for activated (i.e., proliferating) TPPII-KO CD8+ T cells on γ-irradiation (Fig. 1D; Supplementary Fig. S2).

Radiation sensitivity of primary fibroblasts. Fibroblasts are not apoptosis-prone, and we did not find any significant differences in apoptosis between WT and KO fibroblasts on γ-irradiation (Fig. 2A). Nonetheless, we counted more TPPII-deficient fibroblasts with signs of mitotic catastrophe (Fig. 2B and C) but noted that the increase was rather small (although statistically significant).

Figure 2.

Radiation sensitivity of primary fibroblasts. A, primary (passage 2) fibroblasts were irradiated with 5 Gy. Apoptosis was measured by flow cytometry after staining with Annexin V 48 h later. B and C, primary fibroblasts were seeded on coverslips. Twenty-four hours later, cells were irradiated with 5 Gy and, another 48 h later, nuclei were stained with 4′,6-diamidino-2-phenylindole (DAPI). Cells were analyzed on a fluorescence microscope with a 40-fold magnification objective. Two hundred cells per coverslip and five coverslips each were scored. The experiment was done twice.

Figure 2.

Radiation sensitivity of primary fibroblasts. A, primary (passage 2) fibroblasts were irradiated with 5 Gy. Apoptosis was measured by flow cytometry after staining with Annexin V 48 h later. B and C, primary fibroblasts were seeded on coverslips. Twenty-four hours later, cells were irradiated with 5 Gy and, another 48 h later, nuclei were stained with 4′,6-diamidino-2-phenylindole (DAPI). Cells were analyzed on a fluorescence microscope with a 40-fold magnification objective. Two hundred cells per coverslip and five coverslips each were scored. The experiment was done twice.

Close modal

The radiation-induced p53-dependent G1 arrest is functional in TPPII-KO cells. The p53 protein is essential for genotoxic stress–induced G1 cell cycle arrest, during which the damaged DNA should be repaired before cells enter S phase (1820). We assessed the functionality of the G1 checkpoint by flow cytometric analysis of the nuclear DNA content as well as by measuring the incorporation of the nucleotide analogue BrdUrd during S phase on γ-irradiation. As seen in Fig. 3A and B and Supplementary Fig. S3, cell cycle analysis indicated that S phase was strongly reduced in irradiated primary fibroblasts and irradiated activated CD8+ T cells, to a similar extent in WT and KO cells. The combined labeling with DNA dye and BrdUrd revealed an even more pronounced reduction in the proportion of S-phase cells for KO compared with WT, particularly for proliferating CD8+ T cells (Fig. 3C and D).

Figure 3.

Pronounced cell cycle arrest of TPPII-deficient primary fibroblasts and activated T cells on irradiation. A, primary (passage 2) fibroblasts were irradiated with 5 Gy. At the indicated time points thereafter, samples were collected for cell cycle analysis by flow cytometry after staining of fixed cells with propidium iodide. B, isolated spleen and lymph node CD8+ WT and KO T cells were activated in medium supplemented with PMA (10 ng/mL) and ionomycin (500 ng/mL). After 48 h, the cells were irradiated with 5 Gy and samples were collected for cell cycle analysis with propidium iodide at the time points indicated. C, isolated CD8+ WT and KO splenocytes and lymph node cells were activated as described in B. Forty-eight hours later, the cells were irradiated with 5 Gy, and another 8 and 16 h later, BrdUrd (10 μmol/L final concentration) was added for 30 min to label proliferating S-phase cells. Cells were then stained using a BrdUrd staining kit, and BrdUrd incorporation and 7-AAD signal were recorded by flow cytometry. D, summary of the data shown in C. Columns, mean percentage of BrdUrd-positive cells; bars, SD. Each of the experiments in A to D was done three to four times.

Figure 3.

Pronounced cell cycle arrest of TPPII-deficient primary fibroblasts and activated T cells on irradiation. A, primary (passage 2) fibroblasts were irradiated with 5 Gy. At the indicated time points thereafter, samples were collected for cell cycle analysis by flow cytometry after staining of fixed cells with propidium iodide. B, isolated spleen and lymph node CD8+ WT and KO T cells were activated in medium supplemented with PMA (10 ng/mL) and ionomycin (500 ng/mL). After 48 h, the cells were irradiated with 5 Gy and samples were collected for cell cycle analysis with propidium iodide at the time points indicated. C, isolated CD8+ WT and KO splenocytes and lymph node cells were activated as described in B. Forty-eight hours later, the cells were irradiated with 5 Gy, and another 8 and 16 h later, BrdUrd (10 μmol/L final concentration) was added for 30 min to label proliferating S-phase cells. Cells were then stained using a BrdUrd staining kit, and BrdUrd incorporation and 7-AAD signal were recorded by flow cytometry. D, summary of the data shown in C. Columns, mean percentage of BrdUrd-positive cells; bars, SD. Each of the experiments in A to D was done three to four times.

Close modal

Activation of p53 as assessed by Western blot. Finally, we assessed γ-irradiation–induced accumulation and activation of p53 by Western blotting (19, 20). Western blot analyses also did not reveal any significant differences between TPPII-KO and WT cells; p53 accumulation and phosphorylation-mediated activation were essentially indistinguishable for WT and KO for various cell types, including resting and activated lymphoid cells as well as fibroblasts (Fig. 4A–D). In addition, the induction of the cyclin-dependent kinase inhibitor p21, a well-known transcriptional target of p53 and a major regulator of the G1 arrest pathway (19, 20), and of the proapoptotic p53/p73 targets Bax and Fas, which are both important for T-cell apoptosis (21, 22), seemed to be normal in irradiated TPPII-deficient cells (Fig. 4A–D). Why Hong and colleagues (12) observed failure of p53 stabilization in irradiated lymphoma cells stably expressing TPPII-specific siRNA is unclear. One possibility is off-target effects, which are frequently observed in siRNA silencing experiments (23, 24).

Figure 4.

γ-Irradiation efficiently activates and stabilizes p53 and induces known p53/p73 targets in several primary cell types derived from TPPII-KO mice. A, isolated splenic and lymph node B cells were irradiated with 5 Gy directly ex vivo. Samples were collected at the indicated time points and the cellular concentrations of total p53, activated (phospho-Ser15) p53, and p21 proteins were assessed by immunoblotting. Actin levels served as controls. B and C, p53, phospho-p53, and p21 levels in irradiated activated splenic and lymph node CD8+ T cells and primary (passage 2) fibroblasts. CD8+ T cells were first cultured in medium supplemented with PMA (10 ng/mL) and ionomycin (500 ng/mL). At day 3 of activation, the cells were irradiated with 5 Gy, and the samples were collected for immunoblotting at the time points indicated. D, phospho-p53, Bax, and Fas levels in irradiated activated CD8+ T cells. CD8+ T cells were activated with PMA and ionomycin for 4 h and the cells were then irradiated with 8 Gy and the samples were collected at the time points indicated. Representative of experiments done at least thrice independently.

Figure 4.

γ-Irradiation efficiently activates and stabilizes p53 and induces known p53/p73 targets in several primary cell types derived from TPPII-KO mice. A, isolated splenic and lymph node B cells were irradiated with 5 Gy directly ex vivo. Samples were collected at the indicated time points and the cellular concentrations of total p53, activated (phospho-Ser15) p53, and p21 proteins were assessed by immunoblotting. Actin levels served as controls. B and C, p53, phospho-p53, and p21 levels in irradiated activated splenic and lymph node CD8+ T cells and primary (passage 2) fibroblasts. CD8+ T cells were first cultured in medium supplemented with PMA (10 ng/mL) and ionomycin (500 ng/mL). At day 3 of activation, the cells were irradiated with 5 Gy, and the samples were collected for immunoblotting at the time points indicated. D, phospho-p53, Bax, and Fas levels in irradiated activated CD8+ T cells. CD8+ T cells were activated with PMA and ionomycin for 4 h and the cells were then irradiated with 8 Gy and the samples were collected at the time points indicated. Representative of experiments done at least thrice independently.

Close modal

TPPII does not move to the nucleus on γ-irradiation. Although we could not confirm the lack of radiation-induced p53 stabilization in the absence of TPPII, we nevertheless wanted to study other potential DNA damage response alterations. We therefore repeated the experiment described by Hong and colleagues (12) showing translocation of TPPII to the nucleus in EL4 cells within 1 hour after γ-irradiation with 5 Gy. Like Hong and colleagues, we used immunofluorescence microscopy. However, we could not detect any significant nuclear translocation of TPPII on γ-irradiation. Nuclear translocation could also not be detected at other radiation doses, at other time points after irradiation (Fig. 5A and B), or in other cell types such as COS cells or transformed fibroblasts (not shown). To confirm our fluorescence microscopy results, we separated the cytosolic and nuclear fractions by differential centrifugation at different time points after irradiation with 5 Gy and determined TPPII by Western blotting. As seen in Fig. 5B, also with this assay, we could not reveal any significant accumulation of TPPII in the nucleus on γ-irradiation. In contrast, p53 accumulated in the nuclear fraction as expected. Why Hong and colleagues found virtually 100% γ-radiation–induced nuclear translocation of TPPII is unclear to us. The only difference between their experiment and ours lies in the antibody used. Hong and colleagues used a polyclonal chicken antibody raised against purified TPPII protein and we used an affinity-purified rabbit serum against a synthetic peptide corresponding to the NH2 terminus of TPPII. We are convinced of the specificity of our antibody, which detects only one band with the molecular mass of TPPII in Western blots of several types of WT cells (Supplementary Fig. S1), depletes TPPII protein in immunoprecipitation (3), and does not recognize any protein in lysates of TPPII-deficient cells (Supplementary Fig. S1). We confirmed our Western blot results with an anti-TPPII antibody directed against an internal epitope of TPPII (Fig. 5B).

Figure 5.

TPPII does not move to the nucleus on γ-irradiation. A, mouse EL4 lymphoma cells were irradiated with 2, 5, or 10 Gy, and after 1 h, the cells were stained for TPPII (red). Nuclei were counterstained with DAPI (blue). Samples were analyzed by fluorescence microscopy at 100-fold magnification. B, EL-4 cells were irradiated with 5 Gy. At the indicated time points, nuclei and cytosol were separated by differential centrifugation and both fractions were analyzed for TPPII content by Western blotting. Sp1 served as a control for nuclear proteins. p53 accumulated in the nucleus on irradiation as expected.

Figure 5.

TPPII does not move to the nucleus on γ-irradiation. A, mouse EL4 lymphoma cells were irradiated with 2, 5, or 10 Gy, and after 1 h, the cells were stained for TPPII (red). Nuclei were counterstained with DAPI (blue). Samples were analyzed by fluorescence microscopy at 100-fold magnification. B, EL-4 cells were irradiated with 5 Gy. At the indicated time points, nuclei and cytosol were separated by differential centrifugation and both fractions were analyzed for TPPII content by Western blotting. Sp1 served as a control for nuclear proteins. p53 accumulated in the nucleus on irradiation as expected.

Close modal

Analysis of the early DNA double-strand break marker γH2AX. The most frequent type of DNA damage caused by ionizing radiation is DNA double-strand breaks (DSB). The histone H2AX belongs to a cohort of DNA damage response proteins that are phosphorylated by the protein kinases ataxia teleangiectasia mutated (ATM) and Rad3-related protein at the DSB site (20). Phosphorylation on Ser139 yields a form called γH2AX. Although we could not confirm radiation-induced movement of TPPII to the nucleus, it remains nevertheless possible that DSB recognition and repair are affected in the absence of TPPII. There are several cytosolic proteins that can affect the DNA damage response (25, 26). Thus, TPPII may, for example, indirectly affect the DNA damage response through its protease function by altering the concentrations of specific DNA damage response proteins. As seen in Fig. 6A, we indeed observed a delay in the resolution of DNA DSB on γ-irradiation in activated TPPII-KO CD8+ T cells compared with WT cells as judged by the γH2AX signal. This was associated with a stronger ATM Ser1981 phosphorylation in the KO cells. However, we could not find consistent differences in the kinetics of the appearance and decline of γH2AX and phospho-ATM immunoreactivity in resting lymphoid cells (Fig. 6B and C) and other cell types such as testes cells (data not shown) and transformed fibroblasts.1

1

Manuscript in preparation.

The difference between activated KO and WT CD8+ T cells in the decline of the γH2AX signal observed here might therefore be due to the stronger radiation-induced apoptosis of the former (see Fig. 1D) rather than to differences in the DSB repair capacity. Apoptotic DNA damage is known to be associated with DNA DSB (27). Slightly delayed γH2AX signal decline was also found for primary TPPII KO fibroblasts (Fig. 6D), but for fibroblasts the differences between WT and KO were usually not as pronounced as those between WT and KO CD8+ T cells, and the phospho-ATM signals seemed to be very similar between WT and KO. Based on our results on primary fibroblasts and activated CD8+ T cells, at present we cannot rule out subtle differences in DNA damage repair depending on TPPII expression in selected cell types.

Figure 6.

Kinetics of phosphorylation and dephosphorylation of H2AX at Ser139 and of ATM at Ser1981 on irradiation in various TPPII WT and KO cells. The cell types indicated were irradiated with 5 Gy and samples were taken for immunoblot analysis at the time points indicated. Actin served as a control. The cells used were isolated spleen and lymph node CD8+ T cells that had been activated with PMA (10 ng/mL) and ionomycin (500 ng/mL) for 3 d (A), nonactivated B cells (B), nonactivated thymocytes directly ex vivo (C), and primary (passage 2) fibroblasts (D). Representative of experiments done at least thrice independently.

Figure 6.

Kinetics of phosphorylation and dephosphorylation of H2AX at Ser139 and of ATM at Ser1981 on irradiation in various TPPII WT and KO cells. The cell types indicated were irradiated with 5 Gy and samples were taken for immunoblot analysis at the time points indicated. Actin served as a control. The cells used were isolated spleen and lymph node CD8+ T cells that had been activated with PMA (10 ng/mL) and ionomycin (500 ng/mL) for 3 d (A), nonactivated B cells (B), nonactivated thymocytes directly ex vivo (C), and primary (passage 2) fibroblasts (D). Representative of experiments done at least thrice independently.

Close modal

The difference in radiosensitivity between activated WT and TPPII KO CD8+ T cells (see Fig. 1D and Supplementary Fig. S2) raises the possibility that TPPII deficiency might specifically increase the radiosensitivity of proliferating cells, perhaps including tumor cells. However, our unpublished data do not suggest that this can be generalized.2

2

Manuscript in preparation.

Instead, the increased radiation-induced apoptosis of activated TPPII-deficient CD8+ T cells could depend on specific mitogenic and survival pathways operating in activated CD8+ lymphocytes (17).

In conclusion, our data do not confirm a role of TPPII in the DNA damage response based on nuclear TPPII translocation and TPPII-mediated p53 stabilization but nevertheless show increased radiation-induced death of selected nontransformed cell types in the absence of the TPPII protease. Strongly increased cell death was found on γ-irradiation of activated TPPII-deficient CD8+ T cells. Future studies will help to unravel the molecular basis for the increased radiation-induced apoptosis of the activated CD8+ T cells in the absence of TPPII.

No potential conflicts of interest were disclosed.

Note: Supplementary data for this article are available at Cancer Research Online (http://cancerres.aacrjournals.org/).

Grant support: German Research Foundation (NI 368/4-3) and Clotten Foundation (G. Niedermann).

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

We thank Prof. Anca-Ligia Grosu for financial support, Dr. Caro Johner (Max-Planck Institute of Immunobiology, Freiburg, Germany) for providing p53 knockout mice, and Dr. Randy Cassada for critical reading of the manuscript.

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Supplementary data