- Email: [email protected]
Distinct sensitivity of CD8+CD4− and CD8+CD4+ leukemic cell subpopulations to cyclophosphamide and rapamycin in Notch1-induced T-ALL mouse model
Leukemia Research 37 (2013) 1592–1601
Contents lists available at ScienceDirect
Leukemia Research journal homepage: www.elsevier.com/locate/leukres
Distinct sensitivity of CD8+ CD4− and CD8+ CD4+ leukemic cell subpopulations to cyclophosphamide and rapamycin in Notch1-induced T-ALL mouse model Yingchi Zhang a,b , Chunlan Hua a,b , Hui Cheng a,b , Weili Wang a,b , Sha Hao a,b , Jing Xu a,b , Xiaomin Wang a,b , Yingdai Gao a,b , Xiaofan Zhu a,b , Tao Cheng a,b,c,∗ , Weiping Yuan a,b,∗ a State Key Laboratory of Experimental Hematology, Institute of Hematology and Blood Diseases Hospital, Chinese Academy of Medical Sciences & Peking Union Medical College, Tianjin 300020, China b Center for Stem Cell Medicine, Chinese Academy of Medical Sciences, Beijing 100730, China c Department of Radiation Oncology, University of Pittsburgh, School of Medicine, Pittsburgh, PA 15213, USA
a r t i c l e
i n f o
Article history: Received 5 May 2013 Received in revised form 1 July 2013 Accepted 9 September 2013 Available online 18 September 2013 Keywords: T-ALL Rapamycin CTX Cell cycle Apoptosis
a b s t r a c t The Notch1 signaling pathway plays an essential role in cell growth and differentiation. Over-expression of the intracellular Notch1 domain (ICN1) in murine hematopoietic cells is able to induce robust T-cell acute lymphoblastic leukemia (T-ALL) in mice. Here we explored the drug sensitivity of T-ALL cells in two subpopulations of CD8+ CD4+ and CD8+ CD4− cells in Notch1-induced T-ALL mice. We found that Notch1 induced T-ALL cells could be decreased by chemotherapeutic drug cyclophosphamide (CTX). CD8+ CD4− T-ALL cells were more sensitive to CTX treatment than CD8+ CD4+ T-ALL cells. The percentage of apoptotic cells induced by CTX treatment was higher in CD8+ CD4− T-ALL cells. T-ALL cells were also inhibited by inhibitor of mTORC1 rapamycin. CD8+ CD4+ T-ALL cells were more susceptible to rapamycin treatment than CD8+ CD4− T-ALL cells. Rapamycin treatment selectively arrested more CD8+ CD4+ T-ALL cells at G0 phase of cell cycle. A combination of the two drugs significantly improved overall survival of T-ALL bearing mice when compared with CTX or rapamycin alone. These results indicated that CD8+ CD4+ and CD8+ CD4− leukemia cell populations had distinct drug sensitivity. © 2013 Elsevier Ltd. All rights reserved.
1. Introduction T-ALL accounts for 25% of adult ALL and 10–15% of childhood ALL [1–3]. Although the survival rate of T-ALL patients has significantly improved with the advance of therapies, the outcome in adult relapsing T-ALL patients remains poor [4,5]. Several transcription factors, e.g., NOTCH1, LMO1/2, TAL1/2, HOX11, and Ikaros, have been implicated in the initiation and maintenance of TALL [6–9]. Among these, activating mutations of NOTCH1 have been observed in over 50% of T-ALL patients [10]. The NOTCH1 signaling pathway is evolutionarily conserved to regulate T cell growth and differentiation [11]. Notch1 gene encodes a conserved type I trans-membrane receptor, which is activated by the ligands of the Delta/Serrate/Lag-2 family expressed on the surface
∗ Corresponding authors at: State Key Laboratory of Experimental Hematology, Institute of Hematology and Blood Diseases Hospital, Chinese Academy of Medical Sciences & Peking Union Medical College, 288 Nanjing Rd, Tianjin, China. Tel.: +86 22 23909197; fax: +86 22 23902047. E-mail addresses: [email protected] (T. Cheng), [email protected] (W. Yuan). 0145-2126/$ – see front matter © 2013 Elsevier Ltd. All rights reserved. http://dx.doi.org/10.1016/j.leukres.2013.09.007
of neighboring cells. Once activated by ligand, the Notch1 receptors undergo proteolytic cleavage, release ICN1 from the plasma membrane, translocate to the nucleus to stimulate transcription of downstream target genes such as hes1, hey1, c-Myc and cyclin D1 [12–14]. Activating mutation of Notch1 was frequently observed in the human T-ALL cell lines and T-ALL mouse models [15–17]. The potent oncogenicity of activating NOTCH1 mutation has been demonstrated in Notch1-induced T-ALL mouse model. Stable transduction of murine lineage negative (Lin− ) bone marrow (BM) cells with ICN1 can result in T-ALL development with 100% penetrance and this leukemia mouse model has been extensively used in the T-ALL studies [4]. c-MYC has been identified as a direct target gene of NOTCH1 during leukemogenesis [18,19]. PI3K-AKT-mTOR signaling pathway also plays an important role downstream of Notch1 signaling pathway. Inhibition of NOTCH1 signaling pathway suppresses mTOR signaling pathway in T-ALL cell lines [20]. NOTCH1 could facilitate the activation of PI3K/AKT/mTOR signaling pathway by downregulating the expression of PTEN, which in turn is a critical negative regulator of PI3K/AKTmTOR signaling pathway [21]. PTEN posttranslational modification though phosphorylation and oxidation leads to activation of PI3K/AKT/mTOR signaling in T-ALL cells
Y. Zhang et al. / Leukemia Research 37 (2013) 1592–1601
[22]. These results suggest that PI3K-AKT-mTOR signaling pathway plays an important role in T-ALL caused by activating NOTCH1 mutation and provided the rationale for the clinical use of mTOR inhibitors such as rapamycin and its analogs in T-ALL. The chemotherapy regimen hyper-CVAD (fractionated CTX, vincristine, doxorubicin, dexamethasone) is effective for de novo ALL [23–26]. However, many T-ALL patients relapsed due to the development of resistance to CTX and other agents during the course of treatments. The drug resistance is partly due to the heterogeneity of T-ALL cells. The response of drug treatment in different populations of T-ALL cells is unclear. A common feature of heterogeneity to T-ALL is the presence of CD8 and CD4 double positive population (CD8+ CD4+ ) as well as CD8 single positive population (CD8+ CD4− ) in many T-ALL mouse models, including Tal1/Lmo2 T-ALL mouse model, Ikaros-deficient T-ALL mouse model and ICN1 T-ALL mouse model [27–29]. To study the drug sensitivity of heterogeneous T-ALL cells in vivo, we used ICN1 T-ALL mouse model to examine the sensitivity of CD8+ CD4+ and CD8+ CD4− T-ALL cells to the cell-cycle non-specific chemotherapeutic drug CTX and mTORC1 inhibitor rapamycin. We found that CTX induced more apoptosis in CD8+ CD4− T-ALL cells and rapamycin specifically inhibited cell cycle of CD8+ CD4+ T-ALL cells. The results revealed higher sensitivity of CD8+ CD4− leukemia cells to CTX and higher sensitivity of CD8+ CD4+ leukemia cells to rapamycin in a Notch1-induced T-ALL mouse model. We also demonstrated that combined use of CTX and rapamycin is more effective in reducing leukemia cells and prolonging T-ALL mice life span than either agent alone. 2. Materials and methods 2.1. Reagents and antibodies Rapamycin (LC Laboratories) was dissolved in absolute ethanol (Sigma–Aldrich) to derive 10 mg/ml stock solution and stored at −80 ◦ C. For in vivo experiments, rapamycin were prepared from the stock solution daily using sterile phosphate buffer solution (PBS) supplemented with 5% PEG-400 (Sigma–Aldrich) and 5% TWEEN-80 (Sigma–Aldrich). CTX (Sigma–Aldrich) was dissolved in sterile PBS to prepare a 40 mg/ml stock and stored at −80 ◦ C. For in vivo experiments, CTX was diluted in sterile PBS to a final concentration of 4 mg/ml. PE conjugated CD3, PEcy7 conjugated CD4, APC conjugated CD8, PE conjugated Phospho-Akt (Thr308) and PE conjugated Phospho-S6 (Ser244) were from BD Bioscience. PE conjugated Phospho-Akt (Ser473) was from Cell Signaling Technology. 2.2. Mice Wild-type (WT) C57BL/6J mice were obtained from Jackson Laboratory. B6.SJL mice were purchased from Taconic Laboratory. For CTX treatment, 5 × 105 established ICN1 induced T-ALL cells were resuspended in 250 l sterile PBS and transplanted into eight weeks old C57BL/6J mice through the tail vein. Mice were treated with CTX (50 mg/kg) or vehicle only once when the percentage of GFP+ TALL cells in peripheral blood (PB) was greater than 10%. Different dosages of CTX had been tested in leukemia bearing mice and the dose of 50 mg/kg of CTX was chosen for the experiment to obtain suboptimal responses to the CTX treatment. The suboptimal dosage of CTX treatment allows a significant reduction of leukemic cells (instead of higher dosage to kill all leukemic cells) in order to observe and analyze the response of T-ALL cells upon CTX treatment. To test the effect of rapamycin on engraftment of T-ALL cells, 5 × 105 T-ALL cells were transplanted into WT mice. From the next day, these mice were treated with rapamycin at the dose of 1 mg/kg, 4 mg/kg, or 10 mg/kg by I.P. injection once per day for 14 days. To test the effect of rapamycin on proliferation of T-ALL cells, 5 × 105 T-ALL cells were transplanted into WT mice. These mice were treated with rapamycin (4 mg/kg) or vehicle once per day for 3 days or 10 days when the percentage of GFP+ T-ALL cells in PB was greater than 10%. All mice were maintained under a specific pathogen-free facility. All animal protocols were approved by the Institutional Animal Care and Use Committee (IACUC), Institute of Hematology and Blood Disease Hospital, CAMS/PUMC. All surgery was performed under sodium pentobarbital anesthesia, and all efforts were made to minimize suffering. 2.3. Establishment of ICN1 induced T-ALL mouse model Transduction of primary BM cells was performed as previously described [30]. In brief, the plasmids (MSCV-ICN1-IRES-GFP, pKAT and pCMV-VSV-G) were cotransfected into packaging cell line 293T using lipofectamine 2000 (Invitrogen) to generate virus. Virus supernatants were harvested 48 and 72 h after transfection.
1593
Lin− cells from the BM of B6.SJL mice (CD45.1+ ) at the age of 8 weeks were enriched with Lineage Cell Depletion Kit (Miltenyi Biotec) according to the manufacturer’s protocol. The Lin− cells were then transduced with viruses and the transduction efficiency was measured by flow cytometry. Bone marrow nucleated cells (BMNCs, 106 /host) from C57BL/6J mice (CD45.2+ ) at the age of 8 weeks were transplanted into lethally irradiated (9.5 Gy) C57BL/6J mice with 106 ICN1 virus-transduced Lin− cells from B6.SJL mice as indicated in Fig. 1A. Primary CD45.1+ GFP+ T-ALL cells were isolated from BM and spleen of end-stage primary leukemia mice and transplanted into sub-lethal irradiated (4.8 Gy) recipient mice. T-ALL cells were collected from the BM and spleen of the end-stage recipient mice and stored. Collected T-ALL cells were subsequently engrafted in WT mice to induce T-ALL for CTX and rapamycin treatment. 2.4. Flow cytometry analysis Mouse BM single-cell suspension was obtained by flushing ilias, femurs, and tibias as described [31]. GFP+ T-ALL cells in PB, BM and spleen were detected by flow cytometry. To determine the immune-phenotypes of T-ALL cells, BM cells of T-ALL mice were labeled with antibodies against CD4 and CD8. For protein kinase phosphorylation analysis, BM cells of T-ALL mice were labeled with surface antibodies against CD4 and CD8, fixed and permeabilized with BD IntraSure Kit (BD Bioscience), and then intracellularly stained with antibodies against p-S6 (pS244), p-AKT (pT308), or p-AKT (pS473). For cell-cycle analysis, BM cells of T-ALL mice were labeled with CD4 and CD8 antibodies, fixed and permeabilized with BD IntraSure Kit (BD Bioscience), and then intracellularly stained with antibody against PE conjugated Ki67 and Hoechst 33342 (BD Bioscience). For apoptosis assay, BM cells of T-ALL mice labeled with CD4 and CD8 antibodies were stained with PE conjugated Annexin V and 7-AAD (BD Bioscience). Analyses were performed on a LSR II (BD Bioscience). 2.5. RNA extraction and quantitative RT-PCR BM single-cell suspension from T-ALL mice was labeled with antibodies against CD8 and CD4. A total of 5 × 105 CD8+ CD4+ and CD8+ CD4− GFP+ T-ALL cells were sorted with BD FACSAria III (BD Bioscience). Total RNA was extracted with an RNeasy Mini Kit (QIAGEN). Reverse transcription was done using Oligo(dT)18 , 2 × TS Reaction Mix, and TransScript RT/RI Enzyme Mix (Transgen). RT-PCR was done with FastStart Universal SYBR Green Master (Roche), 0.4 M of specific forward and reverse primers, and normalized cDNA. The parameters for the thermal cycling of PCR were as follows: 15 s at 95 ◦ C and 60 s at 60 ◦ C, 40 cycles. All the primer sequences are listed Supplementary Table 1. Supplementary material related to this article can be found, in the online version, at http://dx.doi.org/10.1016/j.leukres.2013.09.007. 2.6. Statistical analysis Student’s t tests were used for comparisons between 2 groups and ANOVA analysis for multiple groups. The Kaplan–Meier method was used to construct survival curves, and survival results were compared using the Mantel–Cox log-rank test. All results represent the average of at least three independent experiments and are expressed as mean ± SD. P < 0.05 was considered to be significantly.
3. Results 3.1. CD8+ CD4+ and CD8+ CD4− T-ALL cells are equi-potent in T-ALL induction in WT mice We established the mouse T-ALL model in which lethally irradiated recipients (C57BL/6J) were transplanted with 106 cells that were transduced with the ICN1 expression viruses (starting with Lin− cells) from B6.SJL mice (CD45.1+ ) and 106 BMNCs from C57BL/6J mice (CD45.2+ ) (Fig. 1A). After 2 to 4 weeks transplantation, PB and BM are dominated by CD45.1+ GFP+ T-ALL cells. We have observed that all the GFP+ T-ALL cells co-expressed CD45.1, indicating that all the GFP+ T-ALL cells were from the donor cells of B6.SJL mice. In the subsequent experiment, we used GFP as the marker to distinguish donor T-ALL cells from the recipient mice. Six weeks after transplantation, gross examination of tissues from the sick mice revealed hepatomegaly, splenomegaly and lymphadenopathy resulting from extensive organ infiltration by lymphoblasts. 100% of the recipient mice that received Notch1overexpressing cells developed T-ALL within 8 weeks. Primary GFP+ T-ALL cells from three end-stage leukemia mice were harvested and then transplanted into sub-lethally recipients for expansion. Transplantation of isolated T-ALL cells into un-treated WT mice
1594
Y. Zhang et al. / Leukemia Research 37 (2013) 1592–1601
Fig. 1. CD8+ CD4+ and CD8+ CD4− T-ALL cells have similar kinetics during the course of T-ALL disease. (A) The T-ALL mouse model. A total of 5 × 105 bone marrow nucleated cells (BMNCs) were transplanted into lethally irradiated recipient mice with 5 × 104 Lin− BMNCs successfully transduced with the MSCV-ICN1-IRES-GFP vector. GFP+ T-ALL cells were isolated from end-stage T-ALL mice by FACS, and transplanted into recipient mice. (B) Surface phenotypes of ICN-1 induced T-ALL cells. BM single-cell suspension of T-ALL mice was labeled with antibodies against CD4 and CD8, and analyzed by flow cytometry. (C) Kaplan–Meier analysis of survival time of T-ALL cells bearing mice. CD8+ CD4+ or CD8+ CD4− T-ALL cells of varying amount were injected into WT mice as indicated (n = 7).
induced T-ALL within 4 weeks with 100% penetrance. To confirm the immunophenotype of T-ALL cells, single-cell suspension from BM of T-ALL mice was labeled with antibodies against CD3, CD4 and CD8 and detected by flow cytometry. CD3 was the common marker of T cell recognition. As expected, all the GFP+ T-ALL cells were CD3 positive, indicating that our leukemia mouse model was T cell leukemia. The results revealed two sub-populations (CD8+ CD4+ and CD8+ CD4− ) in ICN-1 induced T-ALL cells (Fig. 1B). In agreement with previous published results [29], CD8+ CD4+ and CD8+ CD4− T-ALL cells showed similar potency in T-ALL induction upon transplantation into WT mice (Fig. 1C).
3.2. CD8+ CD4+ and CD8+ CD4− T-ALL cells respond differently to CTX treatment CTX is extensively used in treating a variety of cancers including T-ALL [26,32–36]. To investigate the drug resistance of leukemia cells in ICN1 induced T-ALL mouse model, we transplanted 5 × 105 established T-ALL cells into WT recipient mice, and then treated these mice with CTX (50 mg/kg) when the percentage of GFP+ T-ALL cells in PB exceeded 10% (Fig. 2A). Flow cytometry was used to detect the percentage of GFP+ T-ALL cells in PB and BM. Effects of CTX on leukemia cells were examined at 4, 8, 12 and 24 h post treatment. CTX treatment caused substantial loss of leukemia cells in BM starting from 12 h post CTX treatment (Fig. 2B). The loss of spleen weight became apparent after 24 h (Fig. 2C). These results indicated that CTX could reduce T-ALL cells in vivo. Furthermore, we analyzed the response of CD8+ CD4+ and CD8+ CD4− T-ALL cells upon CTX treatment after 12 or 24 h. We found that the percentages of CD8+ CD4+ and CD8+ CD4− T-ALL cells were all reduced in BM after 12 or 24 h CTX treatment when compared with vehicle treatment group (Fig. 2D and E). However, the ratio of CD8+ CD4+ versus CD8+ CD4− T-ALL cells increased at 24 h after CTX treatment (Fig. 2F), indicated that CD8+ CD4− T-ALL cells are more sensitive to CTX than CD8+ CD4+ T-ALL cells.
3.3. CD8+ CD4+ T-ALL cells are more anti-apoptotic upon CTX treatment compared with CD8+ CD4− T-ALL cells CTX could cause tumor repression by activating the intrinsic apoptotic pathway as a result of DNA damage [37]. The next set of experiments examined whether varying sensitivity of CD8+ CD4+ and CD8+ CD4− T-ALL cells to CTX was due to different degree of apoptosis. Single-cell suspension was recovered from BM of leukemia mice at 12 or 24 h after CTX treatment, labeled with ANNEXIN V and 7-AAD, and then analyzed by flow cytometry. CTX exposure led to significant apoptosis of T-ALL cells at both time points but with less effects at 24 h post CTX treatment (Fig. 3A). The percentage of apoptotic cells was comparable in CD8+ CD4+ and CD8+ CD4− leukemia cells at 12 h after CTX treatment. However, apoptosis only occurred in CD8+ CD4− T-ALL cells at 24 h after CTX treatment (Fig. 3B and C). In the next experiment, CD8+ CD4+ and CD8+ CD4− T-ALL cells were sorted from leukemia mice by FACS at 12 and 24 h after CTX or vehicle treatment, respectively. RT-PCR analysis of apoptosis related genes revealed significantly higher expression of anti-apoptosis gene Bcl-2 in CD8+ CD4+ T-ALL cells than that in CD8+ CD4− T-ALL cells at 24 h after CTX treatment (Fig. 3D). In contrast, the expression of pro-apoptosis genes bax, p53 and noxa was lower in CD8+ CD4+ T-ALL cells. These results suggested that different response in gene regulation upon CTX could be the basis of distinct sensitivity of CD8+ CD4+ and CD8+ CD4− T-ALL cells to CTX. 3.4. Rapamycin affects CD8+ CD4+ and CD8+ CD4− leukemic cells differently Phosphorylation states of PI3K/Akt/mTOR signaling pathway in Notch1-induced T-ALL cells were examined by measuring the intracellular phosphorylation levels of p-AKT and p-S6 of T-ALL cells using flow cytometry. To test whether intracellular phosphorylation levels of p-AKT and p-S6 detected by flow cytometry are equivalent to those detected by Western Blot (WB), we firstly examined the phosphorylation levels of p-AKT and p-S6 in Jurkat cells after long-term rapamycin treatment (16 h) using flow cytometry
Y. Zhang et al. / Leukemia Research 37 (2013) 1592–1601
1595
Fig. 2. CD8+ CD4+ and CD8+ CD4− T-ALL cells respond differently to CTX. (A) A schematic representation of the experiment design. WT mice received 5 × 105 T-ALL cells via the tail vein, and treated CTX (50 mg/kg) or vehicle once when percentage of GFP+ T-ALL cells in PB exceeded 10%. (B) Percentage of T-ALL cells in BM of leukemia mice treated with CTX, n = 5. (C) Spleen weight of T-ALL mice at 24 h after CTX, n = 5. (D) Representative flow cytometry. Single-cell suspension was obtained from BM of T-ALL mice at 12 or 24 h after CTX treatment for CD4 and CD8 staining. (E) Percentage of T-ALL cells, CD8− CD4+ T-ALL cells and CD8+ CD4+ T-ALL cells in BM after 12 or 24 h CTX treatment, n = 5. (F) The ratio of CD8+ CD4+ versus CD8+ CD4− T-ALL cells. Data are represented as the mean ± SD, *P < 0.05, **P < 0.01, ***P < 0.001, n = 5.
and WB (Supplementary Figure 1A and B). The results indicated that the result of phosphorylation levels of p-AKT and p-S6 in Jurkat after rapamycin treatment are comparable by both flow cytometry and WB methods. The degree of phosphorylation of S6 and AKT in Notch1-induced T-ALL cells was significantly higher than that in primary thymus CD8+ T cells, but lower than that in T-ALL cell line Jurkat cells (Supplementary Figure 2), confirming that PI3K/AKT/mTOR signaling pathway was active in T-ALL cells. This led us to use rapamycin to treat T-ALL mice. We first examined the effect of rapamycin on engraftment of T-ALL cells in WT mice. Daily vehicle or rapamycin treatment at the dose of 1 mg/kg, 4 mg/kg or 10 mg/kg was initiated on the next day after T-ALL cells injection and lasted for 14 days (Supplementary Figure 3A). Rapamycin inhibited the engraftment of T-ALL cells in mice in a dose-dependent manner (decreasing percentage of GFP+ T-ALL cells in PB and BM and lower spleen weight; Supplementary Figure 3B–E). Rapamycin treatment of T-ALL mice (Fig. 4A) also inhibited the proliferation of T-ALL cells (Fig. 4B and C).
Supplementary material related to this article can be found, in the online version, at http://dx.doi.org/10.1016/j.leukres. 2013.09.007. We then analyzed the sensitivity of CD8+ CD4+ and CD8+ CD4− T-ALL cells upon rapamycin treatment after 3 or 10 days. We found that only the percentage of CD8+ CD4+ T-ALL cells, but not CD8+ CD4− T-ALL cells, was significant lower in BM of leukemia mice after 3 or 10 days rapamycin treatment when compared with vehicle treatment (Fig. 4E). A further analysis of the ratio of CD8+ CD4+ versus CD8+ CD4− T-ALL cells confirmed that CD8+ CD4+ T-ALL cells were more sensitive upon rapmaycin treatment (Fig. 4F). 3.5. Rapamycin arrests more CD8+ CD4+ T-ALL cells into G0 phase in comparison to CD8+ CD4− T-ALL cells Rapamycin treatment did not increase apoptosis of T-ALL cells in BM of leukemia bearing mice (Fig. 5). We then examined the cell cycle of T-ALL cells from leukemia mice treated with rapamycin or
1596
Y. Zhang et al. / Leukemia Research 37 (2013) 1592–1601
Fig. 3. CD8+ CD4− T-ALL cells are more sensitive upon CTX treatment than CD8+ CD4+ T-ALL cells. (A) Apoptosis (Annexin V+ 7-AAD+ cells) induced by CTX. Experiment design was the same as for Fig. 2A. Single-cell suspension was prepared from BM of T-ALL mice at 12 or 24 h after CTX (50 mg/kg) for labeling with Annexin V and 7-AAD. Left panel: representative plot. Right panel: data summary. (B) Single-cell suspension was prepared from BM of T-ALL mice at 12 h after CTX treatment for CD4 and CD8 labeling and Annexin V and 7-AAD staining. (C) Single-cell suspension was prepared from BM of T-ALL mice at 24 h after CTX treatment for CD4 and CD8 labeling and Annexin V and 7-AAD staining. (D) 5 × 105 CD8+ CD4+ and CD8+ CD4− T-ALL cells from BM single-cell suspension of T-ALL mice at 12 or 24 h after CTX treatment (50 mg/kg) were sorted by FACS. Isolated RNA was analyzed by real-time RT PCR, and GAPDH was used as internal control. All experiments were repeated at least 3 times with similar results. Data are represented as the mean ± SD, *P < 0.05, **P < 0.01, ***P < 0.001, n = 5.
Y. Zhang et al. / Leukemia Research 37 (2013) 1592–1601
1597
Fig. 4. CD8+ CD4+ and CD8+ CD4− T-ALL cells respond differently to rapmaycin. (A) A schematic representation of the experiment design. WT mice received 5 × 105 T-ALL cells via the tail vein injection, and were treated rapamycin (4 mg/kg) or vehicle once per day when percentage of GFP+ T-ALL cells in PB exceeded 10%. (B) Percentage of T-ALL cells in PB of leukemia mice after 3 or 10 days vehicle or rapamycin treatment (n = 5). (C) Spleen weight of T-ALL mice at 3 or 10 days rapamycin or vehicle (n = 5). (D) Representative flow cytometry. Single-cell suspension was prepared from BM of leukemia mice treated with vehicle or rapamycin once per day for 3 or 10 days for CD4 and CD8 staining. (E) Percentage of T-ALL cells, CD8− CD4+ T-ALL cells and CD8+ CD4+ T-ALL cells in BM after 3 and 10 days CTX treatment (n = 5). (F) The ratio of CD8+ CD4+ versus CD8+ CD4− T-ALL cells. Data are represented as the mean ± SD, *P < 0.05, **P < 0.01, ***P < 0.001.
vehicle using flow cytometry. The results showed that rapamycin exposure for 3 days but not 10 days arrested more T-ALL cells into G0 phase (Fig. 6A). Because long-term rapamycin treatment leads to the activation of PI3K and AKT through a feedback mechanism [38], we analyzed the AKT phosphorylation of T-ALL cells after 3 and 10 days rapamycin or vehicle treatment. We found that AKT phosphorylation at Thr308 or Ser473 site was not affected by longterm rapamycin treatment (10 days) (Supplementary Figure 4A). In Fig. 4, we had shown that CD8+ CD4+ T-ALL cells were more sensitive to rapamycin. So we speculated the cell cycle status of CD8+ CD4+
and CD8+ CD4− T-ALL cells were different after rapamycin treatment. As expected, we found that rapamycin arrested about 40% CD8+ CD4+ T-ALL cells at G0 phase (vs. 15% in the vehicle control). The percentage of CD8+ CD4− T-ALL cells in G0 phase was not affected by rapamycin (Fig. 6B and C). To confirm the effect of rapamycin on CD8+ CD4+ and CD8+ CD4− T-ALL cells, we analyzed the phosphorylation of mTORC1 target S6 in CD8+ CD4+ and CD8+ CD4− T-ALL cells after 3 or 10 days rapamycin treatment. We found that rapamycin reduced phosphorylation level of S6 in both CD8+ CD4+ and CD8+ CD4− T-ALL cells after 3 or 10 days treatment
1598
Y. Zhang et al. / Leukemia Research 37 (2013) 1592–1601
Fig. 5. Rapamycin treatment doesn’t induce the apoptosis of CD8+ CD4+ and CD8+ CD4− T-ALL cells. Single-cell suspension was prepared from BM of T-ALL mice treated with rapamycin or vehicle for 3 days and 10 days, labeled with antibodies against CD4 and CD8, then stained with Annexin V and 7-AAD (n = 5). (A) Representative plot. (B) Data summary. Data are represented as the mean ± SD, *P < 0.05, **P < 0.01, ***P < 0.001.
(Supplementary Figure 4B), which suggested that rapamycin also inhibited the activity of mTOR1 in CD8+ CD4− T-ALL cells. Supplementary material related to this article can be found, in the online version, at http://dx.doi.org/10.1016/j.leukres. 2013.09.007. Next, we found that the expression of cell cycle kinases CDK2 and CDK4 were lower in CD8+ CD4+ T-ALL cells than in CD8+ CD4− T-ALL sorted from BM of T-ALL mice receiving rapamycin. The expression of cell cycle inhibitor p27 was higher in CD8+ CD4+ leukemia cells (Fig. 6D). These results suggested that the different rapamycin sensitivity of CD8+ CD4+ and CD8+ CD4− T-ALL cells is due to distinct gene expression in response to rapamycin. 3.6. Rapamycin in combination with CTX is more effective on T-ALL mice than rapamycin or CTX alone The results presented above demonstrated that CD8+ CD4− TALL cells are more sensitive to CTX, whereas CD8+ CD4+ T-ALL cells are more susceptible to rapamycin. In the next set of experiments, we found that the CTX/rapamycin combination could more effectively reduce T-ALL cell count (Fig. 7B) and the spleen infiltration by leukemia cells (Fig. 7C) in comparison to either CTX or rapamycin alone. The combination of CTX and rapamycin also significantly prolonged the survival of T-ALL mice. The median survival of vehicle-treated mice was 22.0 days. Rapamycin significantly increased the median survival to 33.5 days. At the single suboptimal dosage of 50 mg/kg as mentioned in method, CTX enhanced survival of leukemic mice with a median survival of 29 days. The combinational uses of rapamycin and CTX significantly enhanced overall survival than treatment with rapamycin or CTX alone with a median survival of 46 days (Fig. 7D and Table 1). 4. Discussion Despite significant improvement in clinical therapy, long-term outcomes for relapsed T-ALL patients remain poor. The relapse may reflect a failure to eliminate leukemia stem cells (LSCs) that retains the ability for self-renewal and differentiation. LSCs have been extensively investigated and well documented in acute myeloid leukemia (AML) [39–41]. The existence of LSCs in TALL is still controversial. In our studies, we have shown that the two subpopulations of T-ALLs (CD8+ CD4+ and CD8+ CD4− ) have similar potency in inducing leukemia in recipient mice
(Fig. 1C), which is consistent with previous report [29]. These results suggest that although ICN1-induced T-ALL cells are heterogeneous, surface markers CD4 and CD8 are not useful in identifying LSCs. Defining the differential response of leukemic cell subpopulations to drug treatment may ultimately lead to the better strategies to maximize the efficacy of chemotherapeutic agents. To clarify the distinct drug sensitivity of heterogeneous T-ALL cells, we compared the response of CD8+ CD4+ and CD8+ CD4− T-ALL cells upon CTX and rapamycin treatment. We found that CD8+ CD4+ T-ALL cells were more resistant to CTX treatment but more sensitive to rapamycin treatment when compared with CD8+ CD4− T-ALL cells. In T cell development, it is known that CD8+ CD4− T-ALL cells are derived from CD8+ CD4+ T-ALL cells. Although neither of these two sup-populations represents LSCs, we speculate that different development stages may be one of the reasons for distinct drug sensitivity of CD8+ CD4+ and CD8+ CD4− T-ALL cells upon CTX and rapmycin treatment. However, whether normal CD8+ CD4+ and CD8+ CD4− T cells have distinct sensitivity response to CTX and rapamycin need to be further investigated. Resistance to chemotherapy is a frequent phenomenon in patients with leukemia. In this study, we demonstrated that although CD8+ CD4+ and CD8+ CD4− T-ALL cells had similar ability to induce leukemia, their sensitivity to the representative chemotherapeutic agent CTX differs. Increased ratio of CD8+ CD4+ versus CD8+ CD4− T-ALL cells at 24 h after CTX treatment indicated that CD8+ CD4− leukemia cells are more sensitive to CTX, possibly due to differences in the anti-apoptotic of CTX. At 24 h after CTX treatment, the expression of anti-apoptotic gene Bcl2 was enhanced while the expression of pro-apoptotic genes Bax, Noxa and p53 was reduced in CD8+ CD4+ leukemia cells when compared with CD8+ CD4− leukemia cells. This finding provided the basis for the drug resistance of CTX in T-ALL patients. mTOR plays an essential role in transmission of proliferative signals from PI3K/AKT signaling pathway and has been explored as a potential therapeutic target in several types of tumors, including hematologic malignancies [42]. mTOR inhibitors, such as rapamycin and its analogs, have shown promising therapeutic effects in preclinical leukemia studies alone or in combination with conventional chemotherapy drugs [43]. However, PI3K and AKT become activated through a feedback mechanism upon longterm rapamycin treatment of some cancer cells [38]. The current
Y. Zhang et al. / Leukemia Research 37 (2013) 1592–1601
1599
Fig. 6. More CD8+ CD4+ T-ALL cells are arrested at the G0 phase after rapamycin treatment. (A) Experiment design was the same as in Fig. 3A. Single-cell suspension was prepared from BM of T-ALL mice treated with rapamycin (4 mg/kg) or vehicle for 3 or 10 days for staining with intracellular antibody against with Ki-67 and hoechst33342. Left panel: representative cell cycle. Right panel: data summary. (B) Single-cell suspension was prepared from BM of T-ALL mice receiving rapamycin or vehicle for 3 days, labeled with antibodies against CD4 and CD8, and then followed by intracellular staining using antibody against with Ki-67 and hoechst33342. (C) Single-cell suspension was prepared from BM of T-ALL mice receiving rapamycin or vehicle for 10 days, labeled with antibodies against CD4 and CD8, and then followed by intracellular staining using antibody against with Ki-67 and hoechst33342. (D) 5 × 105 CD8+ CD4+ and CD8+ CD4− T-ALL cells from BM single-cell suspension of T-ALL mice receiving rapamycin for 3 or 10 days were sorted by FACS. Isolated RNA was analyzed with real-time RT PCR, and GAPDH was used as internal control. All experiments were repeated at least 3 times with similar results. Data are represented as the mean ± SD, *P < 0.05, **P < 0.01, ***P < 0.001, n = 5.
1600
Y. Zhang et al. / Leukemia Research 37 (2013) 1592–1601
Fig. 7. CTX in combination with rapamycin is more effective on T-ALL. (A) A schematic representation of the experiment design. WT mice received 5 × 105 T-ALL cells via the tail vein injection, and were treated CTX (50 mg/kg), rapamycin (4 mg/kg) or both. CTX was administered only once, and rapamycin was given once per day, when percentage of GFP+ T-ALL cells in PB exceeded 10%. (B) GFP+ T-ALL cell flow cytometry. Single-cell suspension was obtained from PB of T-ALL mice treated with vehicle, CTX, rapamycin or both for 3 or 10 days (n = 5). (C) Spleen weight of T-ALL mice 3 or 10 days after vehicle, CTX, rapamycin or both (n = 5). (D) Kaplan–Meier survival curve analysis of T-ALL mice treated with vehicle, CTX, rapamycin or both. All experiments were repeated at least 3 times with similar results (n = 6). Data are represented as the mean ± SD, *P < 0.05, **P < 0.01, ***P < 0.001.
Table 1 Statistical analysis of survival in response to individual treatments and treatment combinations.
P value
Rapa vs vehicle
CTX vs vehicle
Rapa + CTX vs vehicle
Rapa + CTX vs Rapa
Rapa + CTX vs CTX
0.0005
0.017
0.0001
0.0005
0.0005
study did not find the feedback activation of AKT and PI3K in Notch1 induced T-ALL cells upon long-term rapamycin treatment (Supplementary Figure 4A). Instead, CD8+ CD4+ , but not CD8+ CD4− T-ALL cells were specifically inhibited both by short-term (3 days) or long-term (10 days) rapamycin treatment. This probably is one of the reasons for limited effect of rapamycin in treating TALL. Although rapamycin inhibited the activity of mTORC1 in both CD8+ CD4+ and CD8+ CD4− T-ALL cells (Supplementary Figure 4B), we found cell cycle arrest only in CD8+ CD4+ , but not CD8+ CD4− T-ALL cells upon rapamycin treatment. The mechanism of expression changes of cell cycle related genes in CD8+ CD4+ and CD8+ CD4− T-ALL cells after rapamycin treatment need to be further investigated. Our results clearly suggested that the CTX/rapamycin combination could offer added benefits in Notch1-induced T-ALL mouse model. The additive effect is probably based on the preferential apoptotic action of CTX in CD8+ CD4− T-ALL cells and the cell cycle arrest action of rapamycin in CD8+ CD4+ T-ALL cells. Taken together, our results demonstrated previously unappreciated distinct sensitivity of CD8+ CD4+ and CD8+ CD4− Notch1-induced T-ALL cells to chemotherapeutic agents. The strategy of targeting different cell subpopulations may ultimately lead to the development of more
rational treatment for T-ALL and other types of cancer although further studies are needed in human T-ALL patients.
Conflicts of interest The authors declare no conflicts of interest.
Acknowledgements This work was supported by grants from the Ministry of Science and Technology of China (2011CB964801, 2012CB966604, 2011ZX09102-010-04, 2010CB945204, 2013BAI01B09) and from National Natural Science Foundation of China (81090410, 30825017, 81130074, 81070390, 81170470, 81300436). Contributions: Y.Z. carried out the conception, design and performed the experiments, analyzed the data, drafting of the article; C.H., H.C., W.W. and S.H. performed research experiments; J.X. and X.W performed data analysis and interpretation; Y.G. and X.Z. provided critical revisions; T.C. and W.Y. contributed to the concept and design, provided drafting of the article, gave final approval
Y. Zhang et al. / Leukemia Research 37 (2013) 1592–1601
and obtained funding. All authors read and approved the final manuscript. References [1] Goldberg JM, Silverman LB, Levy DE, Dalton VK, Gelber RD, et al. Childhood T-cell acute lymphoblastic leukemia: the Dana-Farber Cancer Institute acute lymphoblastic leukemia consortium experience. J Clin Oncol 2003;21:3616–22. [2] Pui CH, Sandlund JT, Pei D, Campana D, Rivera GK, et al. Improved outcome for children with acute lymphoblastic leukemia: results of Total Therapy Study XIIIB at St Jude Children’s Research Hospital. Blood 2004;104:2690–6. [3] Thiel E, Kranz BR, Raghavachar A, Bartram CR, Loffler H, et al. Prethymic phenotype and genotype of pre-T (CD7+/ER−)-cell leukemia and its clinical significance within adult acute lymphoblastic leukemia. Blood 1989;73:1247–58. [4] Grabher C, von Boehmer H, Look AT. Notch 1 activation in the molecular pathogenesis of T-cell acute lymphoblastic leukaemia. Nat Rev Cancer 2006;6:347–59. [5] Kox C, Zimmermann M, Stanulla M, Leible S, Schrappe M, et al. The favorable effect of activating NOTCH1 receptor mutations on long-term outcome in T-ALL patients treated on the ALL-BFM 2000 protocol can be separated from FBXW7 loss of function. Leukemia 2010;24:2005–13. [6] Armstrong SA, Look AT. Molecular genetics of acute lymphoblastic leukemia. J Clin Oncol 2005;23:6306–15. [7] O’Neil J, Look AT. Mechanisms of transcription factor deregulation in lymphoid cell transformation. Oncogene 2007;26:6838–49. [8] Ferrando AA, Neuberg DS, Staunton J, Loh ML, Huard C, et al. Gene expression signatures define novel oncogenic pathways in T cell acute lymphoblastic leukemia. Cancer Cell 2002;1:75–87. [9] Winandy S, Wu P, Georgopoulos K. A dominant mutation in the Ikaros gene leads to rapid development of leukemia and lymphoma. Cell 1995;83:289–99. [10] Weng AP, Ferrando AA, Lee W, Morris JPT, Silverman LB, et al. Activating mutations of NOTCH1 in human T cell acute lymphoblastic leukemia. Science 2004;306:269–71. [11] Bray SJ. Notch signalling: a simple pathway becomes complex. Nat Rev Mol Cell Biol 2006;7:678–89. [12] Klinakis A, Szabolcs M, Politi K, Kiaris H, Artavanis-Tsakonas S, et al. Myc is a Notch1 transcriptional target and a requisite for Notch1-induced mammary tumorigenesis in mice. Proc Natl Acad Sci U S A 2006;103:9262–7. [13] Ronchini C, Capobianco AJ. Induction of cyclin D1 transcription and CDK2 activity by Notch(ic): implication for cell cycle disruption in transformation by Notch(ic). Mol Cell Biol 2001;21:5925–34. [14] Fischer A, Gessler M. Delta-Notch – and then? Protein interactions and proposed modes of repression by Hes and Hey bHLH factors. Nucleic Acids Res 2007;35:4583–96. [15] Dumortier A, Jeannet R, Kirstetter P, Kleinmann E, Sellars M, et al. Notch activation is an early and critical event during T-Cell leukemogenesis in Ikarosdeficient mice. Mol Cell Biol 2006;26:209–20. [16] Reschly EJ, Spaulding C, Vilimas T, Graham WV, Brumbaugh RL, et al. Notch1 promotes survival of E2A-deficient T cell lymphomas through pre-T cell receptor-dependent and -independent mechanisms. Blood 2006;107:4115–21. [17] O’Neil J, Calvo J, McKenna K, Krishnamoorthy V, Aster JC, et al. Activating Notch1 mutations in mouse models of T-ALL. Blood 2006;107:781–5. [18] Palomero T, Lim WK, Odom DT, Sulis ML, Real PJ, et al. NOTCH1 directly regulates c-MYC and activates a feed-forward-loop transcriptional network promoting leukemic cell growth. Proc Natl Acad Sci U S A 2006;103: 18261–6. [19] Weng AP, Millholland JM, Yashiro-Ohtani Y, Arcangeli ML, Lau A, et al. cMyc is an important direct target of Notch1 in T-cell acute lymphoblastic leukemia/lymphoma. Genes Dev 2006;20:2096–109. [20] Chan SM, Weng AP, Tibshirani R, Aster JC, Utz PJ. Notch signals positively regulate activity of the mTOR pathway in T-cell acute lymphoblastic leukemia. Blood 2007;110:278–86. [21] Palomero T, Sulis ML, Cortina M, Real PJ, Barnes K, et al. Mutational loss of PTEN induces resistance to NOTCH1 inhibition in T-cell leukemia. Nat Med 2007;13:1203–10.
1601
[22] Silva A, Yunes JA, Cardoso BA, Martins LR, Jotta PY, et al. PTEN posttranslational inactivation and hyperactivation of the PI3K/Akt pathway sustain primary T cell leukemia viability. J Clin Invest 2008;118:3762–74. [23] Kantarjian HM, O’Brien S, Smith TL, Cortes J, Giles FJ, et al. Results of treatment with hyper-CVAD, a dose-intensive regimen, in adult acute lymphocytic leukemia. J Clin Oncol 2000;18:547–61. [24] Kantarjian H, Thomas D, O’Brien S, Cortes J, Giles F, et al. Long-term followup results of hyperfractionated cyclophosphamide, vincristine, doxorubicin, and dexamethasone (Hyper-CVAD), a dose-intensive regimen, in adult acute lymphocytic leukemia. Cancer 2004;101:2788–801. [25] Murphy SB, Bowman WP, Abromowitch M, Mirro J, Ochs J, et al. Results of treatment of advanced-stage Burkitt’s lymphoma and B cell (SIg+) acute lymphoblastic leukemia with high-dose fractionated cyclophosphamide and coordinated high-dose methotrexate and cytarabine. J Clin Oncol 1986;4:1732–9. [26] O’Brien S, Thomas DA, Ravandi F, Faderl S, Pierce S, et al. Results of the hyperfractionated cyclophosphamide, vincristine, doxorubicin, and dexamethasone regimen in elderly patients with acute lymphocytic leukemia. Cancer 2008;113:2097–101. [27] Tatarek J, Cullion K, Ashworth T, Gerstein R, Aster JC, et al. Notch1 inhibition targets the leukemia-initiating cells in a Tal1/Lmo2 mouse model of T-ALL. Blood 2011;118:1579–90. [28] Jeannet R, Mastio J, Macias-Garcia A, Oravecz A, Ashworth T, et al. Oncogenic activation of the Notch1 gene by deletion of its promoter in Ikaros-deficient T-ALL. Blood 2010;116:5443–54. [29] Li X, Gounari F, Protopopov A, Khazaie K, von Boehmer H. Oncogenesis of TALL and nonmalignant consequences of overexpressing intracellular NOTCH1. J Exp Med 2008;205:2851–61. [30] Hu X, Shen H, Tian C, Yu H, Zheng G, et al. Kinetics of normal hematopoietic stem and progenitor cells in a Notch1-induced leukemia model. Blood 2009;114:3783–92. [31] Spangrude GJ. Enrichment of murine haemopoietic stem cells: diverging roads. Immunol Today 1989;10:344–50. [32] Grunberg SM. Cyclophosphamide and etoposide for non-small cell and small cell lung cancer. Drugs 1999;58:11–5. Suppl 3. [33] Tabata M, Kiura K, Okimoto N, Segawa Y, Shinkai T, et al. A phase II trial of cisplatin and irinotecan alternating with doxorubicin, cyclophosphamide and etoposide in previously untreated patients with extensive-disease small-cell lung cancer. Cancer Chemother Pharmacol 2007;60:1–6. [34] Nicolini A, Mancini P, Ferrari P, Anselmi L, Tartarelli G, et al. Oral low-dose cyclophosphamide in metastatic hormone refractory prostate cancer (MHRPC). Biomed Pharmacother 2004;58:447–50. [35] Nelius T, Rinard K, Filleur S. Oral/metronomic cyclophosphamide-based chemotherapy as option for patients with castration-refractory prostate cancer: review of the literature. Cancer Treat Rev 2011;37:444–55. [36] Tanaka M, Takamatsu Y, Anan K, Ohno S, Nishimura R, et al. Oral combination chemotherapy with capecitabine and cyclophosphamide in patients with metastatic breast cancer: a phase II study. Anticancer Drugs 2010;21:453–8. [37] Happo L, Cragg MS, Phipson B, Haga JM, Jansen ES, et al. Maximal killing of lymphoma cells by DNA damage-inducing therapy requires not only the p53 targets Puma and Noxa, but also Bim. Blood 2010;116:5256–67. [38] Yap TA, Garrett MD, Walton MI, Raynaud F, de Bono JS, et al. Targeting the PI3KAKT-mTOR pathway: progress, pitfalls, and promises. Curr Opin Pharmacol 2008;8:393–412. [39] Lapidot T, Sirard C, Vormoor J, Murdoch B, Hoang T, et al. A cell initiating human acute myeloid leukaemia after transplantation into SCID mice. Nature 1994;367:645–8. [40] Bonnet D, Dick JE. Human acute myeloid leukemia is organized as a hierarchy that originates from a primitive hematopoietic cell. Nat Med 1997;3:730–7. [41] Krause DS, Van Etten RA. Right on target: eradicating leukemic stem cells. Trends Mol Med 2007;13:470–81. [42] Chiang GG, Abraham RT. Targeting the mTOR signaling network in cancer. Trends Mol Med 2007;13:433–42. [43] Brown VI, Seif AE, Reid GS, Teachey DT, Grupp SA. Novel molecular and cellular therapeutic targets in acute lymphoblastic leukemia and lymphoproliferative disease. Immunol Res 2008;42:84–105.
Contents lists available at ScienceDirect
Leukemia Research journal homepage: www.elsevier.com/locate/leukres
Distinct sensitivity of CD8+ CD4− and CD8+ CD4+ leukemic cell subpopulations to cyclophosphamide and rapamycin in Notch1-induced T-ALL mouse model Yingchi Zhang a,b , Chunlan Hua a,b , Hui Cheng a,b , Weili Wang a,b , Sha Hao a,b , Jing Xu a,b , Xiaomin Wang a,b , Yingdai Gao a,b , Xiaofan Zhu a,b , Tao Cheng a,b,c,∗ , Weiping Yuan a,b,∗ a State Key Laboratory of Experimental Hematology, Institute of Hematology and Blood Diseases Hospital, Chinese Academy of Medical Sciences & Peking Union Medical College, Tianjin 300020, China b Center for Stem Cell Medicine, Chinese Academy of Medical Sciences, Beijing 100730, China c Department of Radiation Oncology, University of Pittsburgh, School of Medicine, Pittsburgh, PA 15213, USA
a r t i c l e
i n f o
Article history: Received 5 May 2013 Received in revised form 1 July 2013 Accepted 9 September 2013 Available online 18 September 2013 Keywords: T-ALL Rapamycin CTX Cell cycle Apoptosis
a b s t r a c t The Notch1 signaling pathway plays an essential role in cell growth and differentiation. Over-expression of the intracellular Notch1 domain (ICN1) in murine hematopoietic cells is able to induce robust T-cell acute lymphoblastic leukemia (T-ALL) in mice. Here we explored the drug sensitivity of T-ALL cells in two subpopulations of CD8+ CD4+ and CD8+ CD4− cells in Notch1-induced T-ALL mice. We found that Notch1 induced T-ALL cells could be decreased by chemotherapeutic drug cyclophosphamide (CTX). CD8+ CD4− T-ALL cells were more sensitive to CTX treatment than CD8+ CD4+ T-ALL cells. The percentage of apoptotic cells induced by CTX treatment was higher in CD8+ CD4− T-ALL cells. T-ALL cells were also inhibited by inhibitor of mTORC1 rapamycin. CD8+ CD4+ T-ALL cells were more susceptible to rapamycin treatment than CD8+ CD4− T-ALL cells. Rapamycin treatment selectively arrested more CD8+ CD4+ T-ALL cells at G0 phase of cell cycle. A combination of the two drugs significantly improved overall survival of T-ALL bearing mice when compared with CTX or rapamycin alone. These results indicated that CD8+ CD4+ and CD8+ CD4− leukemia cell populations had distinct drug sensitivity. © 2013 Elsevier Ltd. All rights reserved.
1. Introduction T-ALL accounts for 25% of adult ALL and 10–15% of childhood ALL [1–3]. Although the survival rate of T-ALL patients has significantly improved with the advance of therapies, the outcome in adult relapsing T-ALL patients remains poor [4,5]. Several transcription factors, e.g., NOTCH1, LMO1/2, TAL1/2, HOX11, and Ikaros, have been implicated in the initiation and maintenance of TALL [6–9]. Among these, activating mutations of NOTCH1 have been observed in over 50% of T-ALL patients [10]. The NOTCH1 signaling pathway is evolutionarily conserved to regulate T cell growth and differentiation [11]. Notch1 gene encodes a conserved type I trans-membrane receptor, which is activated by the ligands of the Delta/Serrate/Lag-2 family expressed on the surface
∗ Corresponding authors at: State Key Laboratory of Experimental Hematology, Institute of Hematology and Blood Diseases Hospital, Chinese Academy of Medical Sciences & Peking Union Medical College, 288 Nanjing Rd, Tianjin, China. Tel.: +86 22 23909197; fax: +86 22 23902047. E-mail addresses: [email protected] (T. Cheng), [email protected] (W. Yuan). 0145-2126/$ – see front matter © 2013 Elsevier Ltd. All rights reserved. http://dx.doi.org/10.1016/j.leukres.2013.09.007
of neighboring cells. Once activated by ligand, the Notch1 receptors undergo proteolytic cleavage, release ICN1 from the plasma membrane, translocate to the nucleus to stimulate transcription of downstream target genes such as hes1, hey1, c-Myc and cyclin D1 [12–14]. Activating mutation of Notch1 was frequently observed in the human T-ALL cell lines and T-ALL mouse models [15–17]. The potent oncogenicity of activating NOTCH1 mutation has been demonstrated in Notch1-induced T-ALL mouse model. Stable transduction of murine lineage negative (Lin− ) bone marrow (BM) cells with ICN1 can result in T-ALL development with 100% penetrance and this leukemia mouse model has been extensively used in the T-ALL studies [4]. c-MYC has been identified as a direct target gene of NOTCH1 during leukemogenesis [18,19]. PI3K-AKT-mTOR signaling pathway also plays an important role downstream of Notch1 signaling pathway. Inhibition of NOTCH1 signaling pathway suppresses mTOR signaling pathway in T-ALL cell lines [20]. NOTCH1 could facilitate the activation of PI3K/AKT/mTOR signaling pathway by downregulating the expression of PTEN, which in turn is a critical negative regulator of PI3K/AKTmTOR signaling pathway [21]. PTEN posttranslational modification though phosphorylation and oxidation leads to activation of PI3K/AKT/mTOR signaling in T-ALL cells
Y. Zhang et al. / Leukemia Research 37 (2013) 1592–1601
[22]. These results suggest that PI3K-AKT-mTOR signaling pathway plays an important role in T-ALL caused by activating NOTCH1 mutation and provided the rationale for the clinical use of mTOR inhibitors such as rapamycin and its analogs in T-ALL. The chemotherapy regimen hyper-CVAD (fractionated CTX, vincristine, doxorubicin, dexamethasone) is effective for de novo ALL [23–26]. However, many T-ALL patients relapsed due to the development of resistance to CTX and other agents during the course of treatments. The drug resistance is partly due to the heterogeneity of T-ALL cells. The response of drug treatment in different populations of T-ALL cells is unclear. A common feature of heterogeneity to T-ALL is the presence of CD8 and CD4 double positive population (CD8+ CD4+ ) as well as CD8 single positive population (CD8+ CD4− ) in many T-ALL mouse models, including Tal1/Lmo2 T-ALL mouse model, Ikaros-deficient T-ALL mouse model and ICN1 T-ALL mouse model [27–29]. To study the drug sensitivity of heterogeneous T-ALL cells in vivo, we used ICN1 T-ALL mouse model to examine the sensitivity of CD8+ CD4+ and CD8+ CD4− T-ALL cells to the cell-cycle non-specific chemotherapeutic drug CTX and mTORC1 inhibitor rapamycin. We found that CTX induced more apoptosis in CD8+ CD4− T-ALL cells and rapamycin specifically inhibited cell cycle of CD8+ CD4+ T-ALL cells. The results revealed higher sensitivity of CD8+ CD4− leukemia cells to CTX and higher sensitivity of CD8+ CD4+ leukemia cells to rapamycin in a Notch1-induced T-ALL mouse model. We also demonstrated that combined use of CTX and rapamycin is more effective in reducing leukemia cells and prolonging T-ALL mice life span than either agent alone. 2. Materials and methods 2.1. Reagents and antibodies Rapamycin (LC Laboratories) was dissolved in absolute ethanol (Sigma–Aldrich) to derive 10 mg/ml stock solution and stored at −80 ◦ C. For in vivo experiments, rapamycin were prepared from the stock solution daily using sterile phosphate buffer solution (PBS) supplemented with 5% PEG-400 (Sigma–Aldrich) and 5% TWEEN-80 (Sigma–Aldrich). CTX (Sigma–Aldrich) was dissolved in sterile PBS to prepare a 40 mg/ml stock and stored at −80 ◦ C. For in vivo experiments, CTX was diluted in sterile PBS to a final concentration of 4 mg/ml. PE conjugated CD3, PEcy7 conjugated CD4, APC conjugated CD8, PE conjugated Phospho-Akt (Thr308) and PE conjugated Phospho-S6 (Ser244) were from BD Bioscience. PE conjugated Phospho-Akt (Ser473) was from Cell Signaling Technology. 2.2. Mice Wild-type (WT) C57BL/6J mice were obtained from Jackson Laboratory. B6.SJL mice were purchased from Taconic Laboratory. For CTX treatment, 5 × 105 established ICN1 induced T-ALL cells were resuspended in 250 l sterile PBS and transplanted into eight weeks old C57BL/6J mice through the tail vein. Mice were treated with CTX (50 mg/kg) or vehicle only once when the percentage of GFP+ TALL cells in peripheral blood (PB) was greater than 10%. Different dosages of CTX had been tested in leukemia bearing mice and the dose of 50 mg/kg of CTX was chosen for the experiment to obtain suboptimal responses to the CTX treatment. The suboptimal dosage of CTX treatment allows a significant reduction of leukemic cells (instead of higher dosage to kill all leukemic cells) in order to observe and analyze the response of T-ALL cells upon CTX treatment. To test the effect of rapamycin on engraftment of T-ALL cells, 5 × 105 T-ALL cells were transplanted into WT mice. From the next day, these mice were treated with rapamycin at the dose of 1 mg/kg, 4 mg/kg, or 10 mg/kg by I.P. injection once per day for 14 days. To test the effect of rapamycin on proliferation of T-ALL cells, 5 × 105 T-ALL cells were transplanted into WT mice. These mice were treated with rapamycin (4 mg/kg) or vehicle once per day for 3 days or 10 days when the percentage of GFP+ T-ALL cells in PB was greater than 10%. All mice were maintained under a specific pathogen-free facility. All animal protocols were approved by the Institutional Animal Care and Use Committee (IACUC), Institute of Hematology and Blood Disease Hospital, CAMS/PUMC. All surgery was performed under sodium pentobarbital anesthesia, and all efforts were made to minimize suffering. 2.3. Establishment of ICN1 induced T-ALL mouse model Transduction of primary BM cells was performed as previously described [30]. In brief, the plasmids (MSCV-ICN1-IRES-GFP, pKAT and pCMV-VSV-G) were cotransfected into packaging cell line 293T using lipofectamine 2000 (Invitrogen) to generate virus. Virus supernatants were harvested 48 and 72 h after transfection.
1593
Lin− cells from the BM of B6.SJL mice (CD45.1+ ) at the age of 8 weeks were enriched with Lineage Cell Depletion Kit (Miltenyi Biotec) according to the manufacturer’s protocol. The Lin− cells were then transduced with viruses and the transduction efficiency was measured by flow cytometry. Bone marrow nucleated cells (BMNCs, 106 /host) from C57BL/6J mice (CD45.2+ ) at the age of 8 weeks were transplanted into lethally irradiated (9.5 Gy) C57BL/6J mice with 106 ICN1 virus-transduced Lin− cells from B6.SJL mice as indicated in Fig. 1A. Primary CD45.1+ GFP+ T-ALL cells were isolated from BM and spleen of end-stage primary leukemia mice and transplanted into sub-lethal irradiated (4.8 Gy) recipient mice. T-ALL cells were collected from the BM and spleen of the end-stage recipient mice and stored. Collected T-ALL cells were subsequently engrafted in WT mice to induce T-ALL for CTX and rapamycin treatment. 2.4. Flow cytometry analysis Mouse BM single-cell suspension was obtained by flushing ilias, femurs, and tibias as described [31]. GFP+ T-ALL cells in PB, BM and spleen were detected by flow cytometry. To determine the immune-phenotypes of T-ALL cells, BM cells of T-ALL mice were labeled with antibodies against CD4 and CD8. For protein kinase phosphorylation analysis, BM cells of T-ALL mice were labeled with surface antibodies against CD4 and CD8, fixed and permeabilized with BD IntraSure Kit (BD Bioscience), and then intracellularly stained with antibodies against p-S6 (pS244), p-AKT (pT308), or p-AKT (pS473). For cell-cycle analysis, BM cells of T-ALL mice were labeled with CD4 and CD8 antibodies, fixed and permeabilized with BD IntraSure Kit (BD Bioscience), and then intracellularly stained with antibody against PE conjugated Ki67 and Hoechst 33342 (BD Bioscience). For apoptosis assay, BM cells of T-ALL mice labeled with CD4 and CD8 antibodies were stained with PE conjugated Annexin V and 7-AAD (BD Bioscience). Analyses were performed on a LSR II (BD Bioscience). 2.5. RNA extraction and quantitative RT-PCR BM single-cell suspension from T-ALL mice was labeled with antibodies against CD8 and CD4. A total of 5 × 105 CD8+ CD4+ and CD8+ CD4− GFP+ T-ALL cells were sorted with BD FACSAria III (BD Bioscience). Total RNA was extracted with an RNeasy Mini Kit (QIAGEN). Reverse transcription was done using Oligo(dT)18 , 2 × TS Reaction Mix, and TransScript RT/RI Enzyme Mix (Transgen). RT-PCR was done with FastStart Universal SYBR Green Master (Roche), 0.4 M of specific forward and reverse primers, and normalized cDNA. The parameters for the thermal cycling of PCR were as follows: 15 s at 95 ◦ C and 60 s at 60 ◦ C, 40 cycles. All the primer sequences are listed Supplementary Table 1. Supplementary material related to this article can be found, in the online version, at http://dx.doi.org/10.1016/j.leukres.2013.09.007. 2.6. Statistical analysis Student’s t tests were used for comparisons between 2 groups and ANOVA analysis for multiple groups. The Kaplan–Meier method was used to construct survival curves, and survival results were compared using the Mantel–Cox log-rank test. All results represent the average of at least three independent experiments and are expressed as mean ± SD. P < 0.05 was considered to be significantly.
3. Results 3.1. CD8+ CD4+ and CD8+ CD4− T-ALL cells are equi-potent in T-ALL induction in WT mice We established the mouse T-ALL model in which lethally irradiated recipients (C57BL/6J) were transplanted with 106 cells that were transduced with the ICN1 expression viruses (starting with Lin− cells) from B6.SJL mice (CD45.1+ ) and 106 BMNCs from C57BL/6J mice (CD45.2+ ) (Fig. 1A). After 2 to 4 weeks transplantation, PB and BM are dominated by CD45.1+ GFP+ T-ALL cells. We have observed that all the GFP+ T-ALL cells co-expressed CD45.1, indicating that all the GFP+ T-ALL cells were from the donor cells of B6.SJL mice. In the subsequent experiment, we used GFP as the marker to distinguish donor T-ALL cells from the recipient mice. Six weeks after transplantation, gross examination of tissues from the sick mice revealed hepatomegaly, splenomegaly and lymphadenopathy resulting from extensive organ infiltration by lymphoblasts. 100% of the recipient mice that received Notch1overexpressing cells developed T-ALL within 8 weeks. Primary GFP+ T-ALL cells from three end-stage leukemia mice were harvested and then transplanted into sub-lethally recipients for expansion. Transplantation of isolated T-ALL cells into un-treated WT mice
1594
Y. Zhang et al. / Leukemia Research 37 (2013) 1592–1601
Fig. 1. CD8+ CD4+ and CD8+ CD4− T-ALL cells have similar kinetics during the course of T-ALL disease. (A) The T-ALL mouse model. A total of 5 × 105 bone marrow nucleated cells (BMNCs) were transplanted into lethally irradiated recipient mice with 5 × 104 Lin− BMNCs successfully transduced with the MSCV-ICN1-IRES-GFP vector. GFP+ T-ALL cells were isolated from end-stage T-ALL mice by FACS, and transplanted into recipient mice. (B) Surface phenotypes of ICN-1 induced T-ALL cells. BM single-cell suspension of T-ALL mice was labeled with antibodies against CD4 and CD8, and analyzed by flow cytometry. (C) Kaplan–Meier analysis of survival time of T-ALL cells bearing mice. CD8+ CD4+ or CD8+ CD4− T-ALL cells of varying amount were injected into WT mice as indicated (n = 7).
induced T-ALL within 4 weeks with 100% penetrance. To confirm the immunophenotype of T-ALL cells, single-cell suspension from BM of T-ALL mice was labeled with antibodies against CD3, CD4 and CD8 and detected by flow cytometry. CD3 was the common marker of T cell recognition. As expected, all the GFP+ T-ALL cells were CD3 positive, indicating that our leukemia mouse model was T cell leukemia. The results revealed two sub-populations (CD8+ CD4+ and CD8+ CD4− ) in ICN-1 induced T-ALL cells (Fig. 1B). In agreement with previous published results [29], CD8+ CD4+ and CD8+ CD4− T-ALL cells showed similar potency in T-ALL induction upon transplantation into WT mice (Fig. 1C).
3.2. CD8+ CD4+ and CD8+ CD4− T-ALL cells respond differently to CTX treatment CTX is extensively used in treating a variety of cancers including T-ALL [26,32–36]. To investigate the drug resistance of leukemia cells in ICN1 induced T-ALL mouse model, we transplanted 5 × 105 established T-ALL cells into WT recipient mice, and then treated these mice with CTX (50 mg/kg) when the percentage of GFP+ T-ALL cells in PB exceeded 10% (Fig. 2A). Flow cytometry was used to detect the percentage of GFP+ T-ALL cells in PB and BM. Effects of CTX on leukemia cells were examined at 4, 8, 12 and 24 h post treatment. CTX treatment caused substantial loss of leukemia cells in BM starting from 12 h post CTX treatment (Fig. 2B). The loss of spleen weight became apparent after 24 h (Fig. 2C). These results indicated that CTX could reduce T-ALL cells in vivo. Furthermore, we analyzed the response of CD8+ CD4+ and CD8+ CD4− T-ALL cells upon CTX treatment after 12 or 24 h. We found that the percentages of CD8+ CD4+ and CD8+ CD4− T-ALL cells were all reduced in BM after 12 or 24 h CTX treatment when compared with vehicle treatment group (Fig. 2D and E). However, the ratio of CD8+ CD4+ versus CD8+ CD4− T-ALL cells increased at 24 h after CTX treatment (Fig. 2F), indicated that CD8+ CD4− T-ALL cells are more sensitive to CTX than CD8+ CD4+ T-ALL cells.
3.3. CD8+ CD4+ T-ALL cells are more anti-apoptotic upon CTX treatment compared with CD8+ CD4− T-ALL cells CTX could cause tumor repression by activating the intrinsic apoptotic pathway as a result of DNA damage [37]. The next set of experiments examined whether varying sensitivity of CD8+ CD4+ and CD8+ CD4− T-ALL cells to CTX was due to different degree of apoptosis. Single-cell suspension was recovered from BM of leukemia mice at 12 or 24 h after CTX treatment, labeled with ANNEXIN V and 7-AAD, and then analyzed by flow cytometry. CTX exposure led to significant apoptosis of T-ALL cells at both time points but with less effects at 24 h post CTX treatment (Fig. 3A). The percentage of apoptotic cells was comparable in CD8+ CD4+ and CD8+ CD4− leukemia cells at 12 h after CTX treatment. However, apoptosis only occurred in CD8+ CD4− T-ALL cells at 24 h after CTX treatment (Fig. 3B and C). In the next experiment, CD8+ CD4+ and CD8+ CD4− T-ALL cells were sorted from leukemia mice by FACS at 12 and 24 h after CTX or vehicle treatment, respectively. RT-PCR analysis of apoptosis related genes revealed significantly higher expression of anti-apoptosis gene Bcl-2 in CD8+ CD4+ T-ALL cells than that in CD8+ CD4− T-ALL cells at 24 h after CTX treatment (Fig. 3D). In contrast, the expression of pro-apoptosis genes bax, p53 and noxa was lower in CD8+ CD4+ T-ALL cells. These results suggested that different response in gene regulation upon CTX could be the basis of distinct sensitivity of CD8+ CD4+ and CD8+ CD4− T-ALL cells to CTX. 3.4. Rapamycin affects CD8+ CD4+ and CD8+ CD4− leukemic cells differently Phosphorylation states of PI3K/Akt/mTOR signaling pathway in Notch1-induced T-ALL cells were examined by measuring the intracellular phosphorylation levels of p-AKT and p-S6 of T-ALL cells using flow cytometry. To test whether intracellular phosphorylation levels of p-AKT and p-S6 detected by flow cytometry are equivalent to those detected by Western Blot (WB), we firstly examined the phosphorylation levels of p-AKT and p-S6 in Jurkat cells after long-term rapamycin treatment (16 h) using flow cytometry
Y. Zhang et al. / Leukemia Research 37 (2013) 1592–1601
1595
Fig. 2. CD8+ CD4+ and CD8+ CD4− T-ALL cells respond differently to CTX. (A) A schematic representation of the experiment design. WT mice received 5 × 105 T-ALL cells via the tail vein, and treated CTX (50 mg/kg) or vehicle once when percentage of GFP+ T-ALL cells in PB exceeded 10%. (B) Percentage of T-ALL cells in BM of leukemia mice treated with CTX, n = 5. (C) Spleen weight of T-ALL mice at 24 h after CTX, n = 5. (D) Representative flow cytometry. Single-cell suspension was obtained from BM of T-ALL mice at 12 or 24 h after CTX treatment for CD4 and CD8 staining. (E) Percentage of T-ALL cells, CD8− CD4+ T-ALL cells and CD8+ CD4+ T-ALL cells in BM after 12 or 24 h CTX treatment, n = 5. (F) The ratio of CD8+ CD4+ versus CD8+ CD4− T-ALL cells. Data are represented as the mean ± SD, *P < 0.05, **P < 0.01, ***P < 0.001, n = 5.
and WB (Supplementary Figure 1A and B). The results indicated that the result of phosphorylation levels of p-AKT and p-S6 in Jurkat after rapamycin treatment are comparable by both flow cytometry and WB methods. The degree of phosphorylation of S6 and AKT in Notch1-induced T-ALL cells was significantly higher than that in primary thymus CD8+ T cells, but lower than that in T-ALL cell line Jurkat cells (Supplementary Figure 2), confirming that PI3K/AKT/mTOR signaling pathway was active in T-ALL cells. This led us to use rapamycin to treat T-ALL mice. We first examined the effect of rapamycin on engraftment of T-ALL cells in WT mice. Daily vehicle or rapamycin treatment at the dose of 1 mg/kg, 4 mg/kg or 10 mg/kg was initiated on the next day after T-ALL cells injection and lasted for 14 days (Supplementary Figure 3A). Rapamycin inhibited the engraftment of T-ALL cells in mice in a dose-dependent manner (decreasing percentage of GFP+ T-ALL cells in PB and BM and lower spleen weight; Supplementary Figure 3B–E). Rapamycin treatment of T-ALL mice (Fig. 4A) also inhibited the proliferation of T-ALL cells (Fig. 4B and C).
Supplementary material related to this article can be found, in the online version, at http://dx.doi.org/10.1016/j.leukres. 2013.09.007. We then analyzed the sensitivity of CD8+ CD4+ and CD8+ CD4− T-ALL cells upon rapamycin treatment after 3 or 10 days. We found that only the percentage of CD8+ CD4+ T-ALL cells, but not CD8+ CD4− T-ALL cells, was significant lower in BM of leukemia mice after 3 or 10 days rapamycin treatment when compared with vehicle treatment (Fig. 4E). A further analysis of the ratio of CD8+ CD4+ versus CD8+ CD4− T-ALL cells confirmed that CD8+ CD4+ T-ALL cells were more sensitive upon rapmaycin treatment (Fig. 4F). 3.5. Rapamycin arrests more CD8+ CD4+ T-ALL cells into G0 phase in comparison to CD8+ CD4− T-ALL cells Rapamycin treatment did not increase apoptosis of T-ALL cells in BM of leukemia bearing mice (Fig. 5). We then examined the cell cycle of T-ALL cells from leukemia mice treated with rapamycin or
1596
Y. Zhang et al. / Leukemia Research 37 (2013) 1592–1601
Fig. 3. CD8+ CD4− T-ALL cells are more sensitive upon CTX treatment than CD8+ CD4+ T-ALL cells. (A) Apoptosis (Annexin V+ 7-AAD+ cells) induced by CTX. Experiment design was the same as for Fig. 2A. Single-cell suspension was prepared from BM of T-ALL mice at 12 or 24 h after CTX (50 mg/kg) for labeling with Annexin V and 7-AAD. Left panel: representative plot. Right panel: data summary. (B) Single-cell suspension was prepared from BM of T-ALL mice at 12 h after CTX treatment for CD4 and CD8 labeling and Annexin V and 7-AAD staining. (C) Single-cell suspension was prepared from BM of T-ALL mice at 24 h after CTX treatment for CD4 and CD8 labeling and Annexin V and 7-AAD staining. (D) 5 × 105 CD8+ CD4+ and CD8+ CD4− T-ALL cells from BM single-cell suspension of T-ALL mice at 12 or 24 h after CTX treatment (50 mg/kg) were sorted by FACS. Isolated RNA was analyzed by real-time RT PCR, and GAPDH was used as internal control. All experiments were repeated at least 3 times with similar results. Data are represented as the mean ± SD, *P < 0.05, **P < 0.01, ***P < 0.001, n = 5.
Y. Zhang et al. / Leukemia Research 37 (2013) 1592–1601
1597
Fig. 4. CD8+ CD4+ and CD8+ CD4− T-ALL cells respond differently to rapmaycin. (A) A schematic representation of the experiment design. WT mice received 5 × 105 T-ALL cells via the tail vein injection, and were treated rapamycin (4 mg/kg) or vehicle once per day when percentage of GFP+ T-ALL cells in PB exceeded 10%. (B) Percentage of T-ALL cells in PB of leukemia mice after 3 or 10 days vehicle or rapamycin treatment (n = 5). (C) Spleen weight of T-ALL mice at 3 or 10 days rapamycin or vehicle (n = 5). (D) Representative flow cytometry. Single-cell suspension was prepared from BM of leukemia mice treated with vehicle or rapamycin once per day for 3 or 10 days for CD4 and CD8 staining. (E) Percentage of T-ALL cells, CD8− CD4+ T-ALL cells and CD8+ CD4+ T-ALL cells in BM after 3 and 10 days CTX treatment (n = 5). (F) The ratio of CD8+ CD4+ versus CD8+ CD4− T-ALL cells. Data are represented as the mean ± SD, *P < 0.05, **P < 0.01, ***P < 0.001.
vehicle using flow cytometry. The results showed that rapamycin exposure for 3 days but not 10 days arrested more T-ALL cells into G0 phase (Fig. 6A). Because long-term rapamycin treatment leads to the activation of PI3K and AKT through a feedback mechanism [38], we analyzed the AKT phosphorylation of T-ALL cells after 3 and 10 days rapamycin or vehicle treatment. We found that AKT phosphorylation at Thr308 or Ser473 site was not affected by longterm rapamycin treatment (10 days) (Supplementary Figure 4A). In Fig. 4, we had shown that CD8+ CD4+ T-ALL cells were more sensitive to rapamycin. So we speculated the cell cycle status of CD8+ CD4+
and CD8+ CD4− T-ALL cells were different after rapamycin treatment. As expected, we found that rapamycin arrested about 40% CD8+ CD4+ T-ALL cells at G0 phase (vs. 15% in the vehicle control). The percentage of CD8+ CD4− T-ALL cells in G0 phase was not affected by rapamycin (Fig. 6B and C). To confirm the effect of rapamycin on CD8+ CD4+ and CD8+ CD4− T-ALL cells, we analyzed the phosphorylation of mTORC1 target S6 in CD8+ CD4+ and CD8+ CD4− T-ALL cells after 3 or 10 days rapamycin treatment. We found that rapamycin reduced phosphorylation level of S6 in both CD8+ CD4+ and CD8+ CD4− T-ALL cells after 3 or 10 days treatment
1598
Y. Zhang et al. / Leukemia Research 37 (2013) 1592–1601
Fig. 5. Rapamycin treatment doesn’t induce the apoptosis of CD8+ CD4+ and CD8+ CD4− T-ALL cells. Single-cell suspension was prepared from BM of T-ALL mice treated with rapamycin or vehicle for 3 days and 10 days, labeled with antibodies against CD4 and CD8, then stained with Annexin V and 7-AAD (n = 5). (A) Representative plot. (B) Data summary. Data are represented as the mean ± SD, *P < 0.05, **P < 0.01, ***P < 0.001.
(Supplementary Figure 4B), which suggested that rapamycin also inhibited the activity of mTOR1 in CD8+ CD4− T-ALL cells. Supplementary material related to this article can be found, in the online version, at http://dx.doi.org/10.1016/j.leukres. 2013.09.007. Next, we found that the expression of cell cycle kinases CDK2 and CDK4 were lower in CD8+ CD4+ T-ALL cells than in CD8+ CD4− T-ALL sorted from BM of T-ALL mice receiving rapamycin. The expression of cell cycle inhibitor p27 was higher in CD8+ CD4+ leukemia cells (Fig. 6D). These results suggested that the different rapamycin sensitivity of CD8+ CD4+ and CD8+ CD4− T-ALL cells is due to distinct gene expression in response to rapamycin. 3.6. Rapamycin in combination with CTX is more effective on T-ALL mice than rapamycin or CTX alone The results presented above demonstrated that CD8+ CD4− TALL cells are more sensitive to CTX, whereas CD8+ CD4+ T-ALL cells are more susceptible to rapamycin. In the next set of experiments, we found that the CTX/rapamycin combination could more effectively reduce T-ALL cell count (Fig. 7B) and the spleen infiltration by leukemia cells (Fig. 7C) in comparison to either CTX or rapamycin alone. The combination of CTX and rapamycin also significantly prolonged the survival of T-ALL mice. The median survival of vehicle-treated mice was 22.0 days. Rapamycin significantly increased the median survival to 33.5 days. At the single suboptimal dosage of 50 mg/kg as mentioned in method, CTX enhanced survival of leukemic mice with a median survival of 29 days. The combinational uses of rapamycin and CTX significantly enhanced overall survival than treatment with rapamycin or CTX alone with a median survival of 46 days (Fig. 7D and Table 1). 4. Discussion Despite significant improvement in clinical therapy, long-term outcomes for relapsed T-ALL patients remain poor. The relapse may reflect a failure to eliminate leukemia stem cells (LSCs) that retains the ability for self-renewal and differentiation. LSCs have been extensively investigated and well documented in acute myeloid leukemia (AML) [39–41]. The existence of LSCs in TALL is still controversial. In our studies, we have shown that the two subpopulations of T-ALLs (CD8+ CD4+ and CD8+ CD4− ) have similar potency in inducing leukemia in recipient mice
(Fig. 1C), which is consistent with previous report [29]. These results suggest that although ICN1-induced T-ALL cells are heterogeneous, surface markers CD4 and CD8 are not useful in identifying LSCs. Defining the differential response of leukemic cell subpopulations to drug treatment may ultimately lead to the better strategies to maximize the efficacy of chemotherapeutic agents. To clarify the distinct drug sensitivity of heterogeneous T-ALL cells, we compared the response of CD8+ CD4+ and CD8+ CD4− T-ALL cells upon CTX and rapamycin treatment. We found that CD8+ CD4+ T-ALL cells were more resistant to CTX treatment but more sensitive to rapamycin treatment when compared with CD8+ CD4− T-ALL cells. In T cell development, it is known that CD8+ CD4− T-ALL cells are derived from CD8+ CD4+ T-ALL cells. Although neither of these two sup-populations represents LSCs, we speculate that different development stages may be one of the reasons for distinct drug sensitivity of CD8+ CD4+ and CD8+ CD4− T-ALL cells upon CTX and rapmycin treatment. However, whether normal CD8+ CD4+ and CD8+ CD4− T cells have distinct sensitivity response to CTX and rapamycin need to be further investigated. Resistance to chemotherapy is a frequent phenomenon in patients with leukemia. In this study, we demonstrated that although CD8+ CD4+ and CD8+ CD4− T-ALL cells had similar ability to induce leukemia, their sensitivity to the representative chemotherapeutic agent CTX differs. Increased ratio of CD8+ CD4+ versus CD8+ CD4− T-ALL cells at 24 h after CTX treatment indicated that CD8+ CD4− leukemia cells are more sensitive to CTX, possibly due to differences in the anti-apoptotic of CTX. At 24 h after CTX treatment, the expression of anti-apoptotic gene Bcl2 was enhanced while the expression of pro-apoptotic genes Bax, Noxa and p53 was reduced in CD8+ CD4+ leukemia cells when compared with CD8+ CD4− leukemia cells. This finding provided the basis for the drug resistance of CTX in T-ALL patients. mTOR plays an essential role in transmission of proliferative signals from PI3K/AKT signaling pathway and has been explored as a potential therapeutic target in several types of tumors, including hematologic malignancies [42]. mTOR inhibitors, such as rapamycin and its analogs, have shown promising therapeutic effects in preclinical leukemia studies alone or in combination with conventional chemotherapy drugs [43]. However, PI3K and AKT become activated through a feedback mechanism upon longterm rapamycin treatment of some cancer cells [38]. The current
Y. Zhang et al. / Leukemia Research 37 (2013) 1592–1601
1599
Fig. 6. More CD8+ CD4+ T-ALL cells are arrested at the G0 phase after rapamycin treatment. (A) Experiment design was the same as in Fig. 3A. Single-cell suspension was prepared from BM of T-ALL mice treated with rapamycin (4 mg/kg) or vehicle for 3 or 10 days for staining with intracellular antibody against with Ki-67 and hoechst33342. Left panel: representative cell cycle. Right panel: data summary. (B) Single-cell suspension was prepared from BM of T-ALL mice receiving rapamycin or vehicle for 3 days, labeled with antibodies against CD4 and CD8, and then followed by intracellular staining using antibody against with Ki-67 and hoechst33342. (C) Single-cell suspension was prepared from BM of T-ALL mice receiving rapamycin or vehicle for 10 days, labeled with antibodies against CD4 and CD8, and then followed by intracellular staining using antibody against with Ki-67 and hoechst33342. (D) 5 × 105 CD8+ CD4+ and CD8+ CD4− T-ALL cells from BM single-cell suspension of T-ALL mice receiving rapamycin for 3 or 10 days were sorted by FACS. Isolated RNA was analyzed with real-time RT PCR, and GAPDH was used as internal control. All experiments were repeated at least 3 times with similar results. Data are represented as the mean ± SD, *P < 0.05, **P < 0.01, ***P < 0.001, n = 5.
1600
Y. Zhang et al. / Leukemia Research 37 (2013) 1592–1601
Fig. 7. CTX in combination with rapamycin is more effective on T-ALL. (A) A schematic representation of the experiment design. WT mice received 5 × 105 T-ALL cells via the tail vein injection, and were treated CTX (50 mg/kg), rapamycin (4 mg/kg) or both. CTX was administered only once, and rapamycin was given once per day, when percentage of GFP+ T-ALL cells in PB exceeded 10%. (B) GFP+ T-ALL cell flow cytometry. Single-cell suspension was obtained from PB of T-ALL mice treated with vehicle, CTX, rapamycin or both for 3 or 10 days (n = 5). (C) Spleen weight of T-ALL mice 3 or 10 days after vehicle, CTX, rapamycin or both (n = 5). (D) Kaplan–Meier survival curve analysis of T-ALL mice treated with vehicle, CTX, rapamycin or both. All experiments were repeated at least 3 times with similar results (n = 6). Data are represented as the mean ± SD, *P < 0.05, **P < 0.01, ***P < 0.001.
Table 1 Statistical analysis of survival in response to individual treatments and treatment combinations.
P value
Rapa vs vehicle
CTX vs vehicle
Rapa + CTX vs vehicle
Rapa + CTX vs Rapa
Rapa + CTX vs CTX
0.0005
0.017
0.0001
0.0005
0.0005
study did not find the feedback activation of AKT and PI3K in Notch1 induced T-ALL cells upon long-term rapamycin treatment (Supplementary Figure 4A). Instead, CD8+ CD4+ , but not CD8+ CD4− T-ALL cells were specifically inhibited both by short-term (3 days) or long-term (10 days) rapamycin treatment. This probably is one of the reasons for limited effect of rapamycin in treating TALL. Although rapamycin inhibited the activity of mTORC1 in both CD8+ CD4+ and CD8+ CD4− T-ALL cells (Supplementary Figure 4B), we found cell cycle arrest only in CD8+ CD4+ , but not CD8+ CD4− T-ALL cells upon rapamycin treatment. The mechanism of expression changes of cell cycle related genes in CD8+ CD4+ and CD8+ CD4− T-ALL cells after rapamycin treatment need to be further investigated. Our results clearly suggested that the CTX/rapamycin combination could offer added benefits in Notch1-induced T-ALL mouse model. The additive effect is probably based on the preferential apoptotic action of CTX in CD8+ CD4− T-ALL cells and the cell cycle arrest action of rapamycin in CD8+ CD4+ T-ALL cells. Taken together, our results demonstrated previously unappreciated distinct sensitivity of CD8+ CD4+ and CD8+ CD4− Notch1-induced T-ALL cells to chemotherapeutic agents. The strategy of targeting different cell subpopulations may ultimately lead to the development of more
rational treatment for T-ALL and other types of cancer although further studies are needed in human T-ALL patients.
Conflicts of interest The authors declare no conflicts of interest.
Acknowledgements This work was supported by grants from the Ministry of Science and Technology of China (2011CB964801, 2012CB966604, 2011ZX09102-010-04, 2010CB945204, 2013BAI01B09) and from National Natural Science Foundation of China (81090410, 30825017, 81130074, 81070390, 81170470, 81300436). Contributions: Y.Z. carried out the conception, design and performed the experiments, analyzed the data, drafting of the article; C.H., H.C., W.W. and S.H. performed research experiments; J.X. and X.W performed data analysis and interpretation; Y.G. and X.Z. provided critical revisions; T.C. and W.Y. contributed to the concept and design, provided drafting of the article, gave final approval
Y. Zhang et al. / Leukemia Research 37 (2013) 1592–1601
and obtained funding. All authors read and approved the final manuscript. References [1] Goldberg JM, Silverman LB, Levy DE, Dalton VK, Gelber RD, et al. Childhood T-cell acute lymphoblastic leukemia: the Dana-Farber Cancer Institute acute lymphoblastic leukemia consortium experience. J Clin Oncol 2003;21:3616–22. [2] Pui CH, Sandlund JT, Pei D, Campana D, Rivera GK, et al. Improved outcome for children with acute lymphoblastic leukemia: results of Total Therapy Study XIIIB at St Jude Children’s Research Hospital. Blood 2004;104:2690–6. [3] Thiel E, Kranz BR, Raghavachar A, Bartram CR, Loffler H, et al. Prethymic phenotype and genotype of pre-T (CD7+/ER−)-cell leukemia and its clinical significance within adult acute lymphoblastic leukemia. Blood 1989;73:1247–58. [4] Grabher C, von Boehmer H, Look AT. Notch 1 activation in the molecular pathogenesis of T-cell acute lymphoblastic leukaemia. Nat Rev Cancer 2006;6:347–59. [5] Kox C, Zimmermann M, Stanulla M, Leible S, Schrappe M, et al. The favorable effect of activating NOTCH1 receptor mutations on long-term outcome in T-ALL patients treated on the ALL-BFM 2000 protocol can be separated from FBXW7 loss of function. Leukemia 2010;24:2005–13. [6] Armstrong SA, Look AT. Molecular genetics of acute lymphoblastic leukemia. J Clin Oncol 2005;23:6306–15. [7] O’Neil J, Look AT. Mechanisms of transcription factor deregulation in lymphoid cell transformation. Oncogene 2007;26:6838–49. [8] Ferrando AA, Neuberg DS, Staunton J, Loh ML, Huard C, et al. Gene expression signatures define novel oncogenic pathways in T cell acute lymphoblastic leukemia. Cancer Cell 2002;1:75–87. [9] Winandy S, Wu P, Georgopoulos K. A dominant mutation in the Ikaros gene leads to rapid development of leukemia and lymphoma. Cell 1995;83:289–99. [10] Weng AP, Ferrando AA, Lee W, Morris JPT, Silverman LB, et al. Activating mutations of NOTCH1 in human T cell acute lymphoblastic leukemia. Science 2004;306:269–71. [11] Bray SJ. Notch signalling: a simple pathway becomes complex. Nat Rev Mol Cell Biol 2006;7:678–89. [12] Klinakis A, Szabolcs M, Politi K, Kiaris H, Artavanis-Tsakonas S, et al. Myc is a Notch1 transcriptional target and a requisite for Notch1-induced mammary tumorigenesis in mice. Proc Natl Acad Sci U S A 2006;103:9262–7. [13] Ronchini C, Capobianco AJ. Induction of cyclin D1 transcription and CDK2 activity by Notch(ic): implication for cell cycle disruption in transformation by Notch(ic). Mol Cell Biol 2001;21:5925–34. [14] Fischer A, Gessler M. Delta-Notch – and then? Protein interactions and proposed modes of repression by Hes and Hey bHLH factors. Nucleic Acids Res 2007;35:4583–96. [15] Dumortier A, Jeannet R, Kirstetter P, Kleinmann E, Sellars M, et al. Notch activation is an early and critical event during T-Cell leukemogenesis in Ikarosdeficient mice. Mol Cell Biol 2006;26:209–20. [16] Reschly EJ, Spaulding C, Vilimas T, Graham WV, Brumbaugh RL, et al. Notch1 promotes survival of E2A-deficient T cell lymphomas through pre-T cell receptor-dependent and -independent mechanisms. Blood 2006;107:4115–21. [17] O’Neil J, Calvo J, McKenna K, Krishnamoorthy V, Aster JC, et al. Activating Notch1 mutations in mouse models of T-ALL. Blood 2006;107:781–5. [18] Palomero T, Lim WK, Odom DT, Sulis ML, Real PJ, et al. NOTCH1 directly regulates c-MYC and activates a feed-forward-loop transcriptional network promoting leukemic cell growth. Proc Natl Acad Sci U S A 2006;103: 18261–6. [19] Weng AP, Millholland JM, Yashiro-Ohtani Y, Arcangeli ML, Lau A, et al. cMyc is an important direct target of Notch1 in T-cell acute lymphoblastic leukemia/lymphoma. Genes Dev 2006;20:2096–109. [20] Chan SM, Weng AP, Tibshirani R, Aster JC, Utz PJ. Notch signals positively regulate activity of the mTOR pathway in T-cell acute lymphoblastic leukemia. Blood 2007;110:278–86. [21] Palomero T, Sulis ML, Cortina M, Real PJ, Barnes K, et al. Mutational loss of PTEN induces resistance to NOTCH1 inhibition in T-cell leukemia. Nat Med 2007;13:1203–10.
1601
[22] Silva A, Yunes JA, Cardoso BA, Martins LR, Jotta PY, et al. PTEN posttranslational inactivation and hyperactivation of the PI3K/Akt pathway sustain primary T cell leukemia viability. J Clin Invest 2008;118:3762–74. [23] Kantarjian HM, O’Brien S, Smith TL, Cortes J, Giles FJ, et al. Results of treatment with hyper-CVAD, a dose-intensive regimen, in adult acute lymphocytic leukemia. J Clin Oncol 2000;18:547–61. [24] Kantarjian H, Thomas D, O’Brien S, Cortes J, Giles F, et al. Long-term followup results of hyperfractionated cyclophosphamide, vincristine, doxorubicin, and dexamethasone (Hyper-CVAD), a dose-intensive regimen, in adult acute lymphocytic leukemia. Cancer 2004;101:2788–801. [25] Murphy SB, Bowman WP, Abromowitch M, Mirro J, Ochs J, et al. Results of treatment of advanced-stage Burkitt’s lymphoma and B cell (SIg+) acute lymphoblastic leukemia with high-dose fractionated cyclophosphamide and coordinated high-dose methotrexate and cytarabine. J Clin Oncol 1986;4:1732–9. [26] O’Brien S, Thomas DA, Ravandi F, Faderl S, Pierce S, et al. Results of the hyperfractionated cyclophosphamide, vincristine, doxorubicin, and dexamethasone regimen in elderly patients with acute lymphocytic leukemia. Cancer 2008;113:2097–101. [27] Tatarek J, Cullion K, Ashworth T, Gerstein R, Aster JC, et al. Notch1 inhibition targets the leukemia-initiating cells in a Tal1/Lmo2 mouse model of T-ALL. Blood 2011;118:1579–90. [28] Jeannet R, Mastio J, Macias-Garcia A, Oravecz A, Ashworth T, et al. Oncogenic activation of the Notch1 gene by deletion of its promoter in Ikaros-deficient T-ALL. Blood 2010;116:5443–54. [29] Li X, Gounari F, Protopopov A, Khazaie K, von Boehmer H. Oncogenesis of TALL and nonmalignant consequences of overexpressing intracellular NOTCH1. J Exp Med 2008;205:2851–61. [30] Hu X, Shen H, Tian C, Yu H, Zheng G, et al. Kinetics of normal hematopoietic stem and progenitor cells in a Notch1-induced leukemia model. Blood 2009;114:3783–92. [31] Spangrude GJ. Enrichment of murine haemopoietic stem cells: diverging roads. Immunol Today 1989;10:344–50. [32] Grunberg SM. Cyclophosphamide and etoposide for non-small cell and small cell lung cancer. Drugs 1999;58:11–5. Suppl 3. [33] Tabata M, Kiura K, Okimoto N, Segawa Y, Shinkai T, et al. A phase II trial of cisplatin and irinotecan alternating with doxorubicin, cyclophosphamide and etoposide in previously untreated patients with extensive-disease small-cell lung cancer. Cancer Chemother Pharmacol 2007;60:1–6. [34] Nicolini A, Mancini P, Ferrari P, Anselmi L, Tartarelli G, et al. Oral low-dose cyclophosphamide in metastatic hormone refractory prostate cancer (MHRPC). Biomed Pharmacother 2004;58:447–50. [35] Nelius T, Rinard K, Filleur S. Oral/metronomic cyclophosphamide-based chemotherapy as option for patients with castration-refractory prostate cancer: review of the literature. Cancer Treat Rev 2011;37:444–55. [36] Tanaka M, Takamatsu Y, Anan K, Ohno S, Nishimura R, et al. Oral combination chemotherapy with capecitabine and cyclophosphamide in patients with metastatic breast cancer: a phase II study. Anticancer Drugs 2010;21:453–8. [37] Happo L, Cragg MS, Phipson B, Haga JM, Jansen ES, et al. Maximal killing of lymphoma cells by DNA damage-inducing therapy requires not only the p53 targets Puma and Noxa, but also Bim. Blood 2010;116:5256–67. [38] Yap TA, Garrett MD, Walton MI, Raynaud F, de Bono JS, et al. Targeting the PI3KAKT-mTOR pathway: progress, pitfalls, and promises. Curr Opin Pharmacol 2008;8:393–412. [39] Lapidot T, Sirard C, Vormoor J, Murdoch B, Hoang T, et al. A cell initiating human acute myeloid leukaemia after transplantation into SCID mice. Nature 1994;367:645–8. [40] Bonnet D, Dick JE. Human acute myeloid leukemia is organized as a hierarchy that originates from a primitive hematopoietic cell. Nat Med 1997;3:730–7. [41] Krause DS, Van Etten RA. Right on target: eradicating leukemic stem cells. Trends Mol Med 2007;13:470–81. [42] Chiang GG, Abraham RT. Targeting the mTOR signaling network in cancer. Trends Mol Med 2007;13:433–42. [43] Brown VI, Seif AE, Reid GS, Teachey DT, Grupp SA. Novel molecular and cellular therapeutic targets in acute lymphoblastic leukemia and lymphoproliferative disease. Immunol Res 2008;42:84–105.