The first two authors contributed equally to this work.
Cancer Cell Biology
Fluorescence lifetime imaging microscopy of chemotherapy-induced apoptosis resistance in a syngenic mouse tumor model
Version of Record online: 8 JUL 2009
Copyright © 2009 UICC
International Journal of Cancer
Volume 126, Issue 1, pages 104–113, 1 January 2010
How to Cite
Keese, M., Yagublu, V., Schwenke, K., Post, S. and Bastiaens, P. (2010), Fluorescence lifetime imaging microscopy of chemotherapy-induced apoptosis resistance in a syngenic mouse tumor model. Int. J. Cancer, 126: 104–113. doi: 10.1002/ijc.24730
- Issue online: 11 NOV 2009
- Version of Record online: 8 JUL 2009
- Accepted manuscript online: 8 JUL 2009 12:00AM EST
- Manuscript Accepted: 23 JUN 2009
- Manuscript Received: 27 JAN 2009
- fluorescence lifetime imaging microscopy (FLIM);
- colon carcinoma;
During cancer therapy with DNA-damaging drug-agents, the development of secondary resistance to apoptosis can be observed. In the search for novel therapeutic approaches that can be used in these cases, we monitored chemotherapy-induced apoptosis resistance in a syngenic mouse tumor model. For this, syngenic murine colorectal carcinoma cells, which stably expressed a FRET-based caspase-3 activity sensor, were introduced into animals to induce peritoneal carcinomatosis or disseminated hepatic metastases. This syngenic system allowed in vitro, in vivo and ex vivo analysis of chemotherapy induced apoptosis induction by optically monitoring the caspase-3 sensor state in the tumor cells. Tumor tissue analysis of 5-FU treated mice showed the selection of 5-FU-induced apoptosis resistant tumor cells. These and chemo-naive fluorescent tumor cells could be re-isolated from treated and untreated mice and propagated in cell culture. Re-exposure to 5-FU and second line treatment modalities in this ex-vivo setting showed that 5-FU induced apoptosis resistance could be alleviated by imatinib mesylate (Gleevec). We thus show that syngenic mouse systems that stably express a FRET-based caspase-3 sensor can be employed to analyse the therapeutic efficiency of apoptosis inducing chemotherapy.
DNA-damaging compounds such as 5-fluorouracil (FU) induce apoptosis by affecting DNA synthesis and repair mechanisms.1 The response to DNA damage is either repair or cell death, and therefore results clinically in chemo-resistance or tumor chemosensitivity.2 The capacity of a cancer cell to repair DNA essentially determines the resistance to chemotherapeutic drugs that induce DNA damage.3 Several signaling pathways have been shown to arrest the cell cycle following DNA damage to allow time for DNA repair. When repair remains incomplete, i.e., in cases of excessive DNA damage, cells will still undergo apoptosis or go into senescence.4 Chemoresistance may therefore be defined as failure of tumor cells to undergo apoptosis in the presence of DNA-damaging compounds. Tumors may be either intrinsically resistant to chemotherapy prior to treatment or acquire resistance during treatment after initial sensitivity.2, 3
The possibility to investigate cancer experimentally in rodent models has provided insights into substeps and mechanisms of disease propagation and guided efforts towards novel cancer therapies. We have previously established a syngenic murine model for colorectal cancer using EGFP as marker protein to determine the tumor load within Balb/c mice.5 The murine C-26 cells used in this model have been described to undergo apoptosis under chemotherapy which implies that they are not primarily resistant to cytotoxic treatment.6 Similar syngenic tumor models have also been developed for squamous cell lung cancer, protastatic cancer and malignant melanoma.7–9 While these models allow evaluation of early tumor development and chemotherapy treatment efficiency by in vivo detection of cells,5 they do not provide any information on the molecular mechanisms affected by chemotherapy. To better understand the molecular background of tumor development such as chemotherapy induced apoptosis resistance, it is necessary to observe key molecular mechanisms which are involved in apoptosis induction in vivo. This could allow the development of novel therapies by facilitating direct monitoring of the influence of chemotherapeutic drugs on molecular reactions that determine the fate of the cell.
Optical imaging of specific molecular targets and pathways in living cells has recently become possible through continued developments in microscopic imaging technology, and more importantly, the availability of genetically encoded fluorescent biosensors.10, 11 These fluorescent reporters have revolutionised live-cell imaging of biochemical processes in a variety of cells from different organisms12–14 since these proteins can directly be used to study protein reactions in living cells by fluorescence resonance energy transfer (FRET).15, 16 Accordingly, FRET between tandems of fluorescent protein variants has been extensively used in living cells for calcium sensing,17, 18 detection of protein interactions,19 detection of enzymatic activities20–22 and monitoring conformational transitions.23–25 Fusion of two fluorescent proteins connected by a short caspase-specific recognition sequence has been also successfully used as an activity-indicator of intracellular caspases – mediators of apoptosis.26–28 The expression of such fusion proteins in transgenic mice has so far been problematic and up to now no genetically encoded FRET-based sensor has been successfully employed in a tumor animal model.
To close this gap, we here introduce a FRET-based method to quantitatively image apoptosis via caspase-3 induced sensor cleavage within individual tumor cells in living mice. Such a system should allow the evaluation of the efficiency of chemotherapy-induced apoptosis of tumor cells within an animal model.
Material and Methods
Primers were synthesized by Thermo Electron (Ulm, Germany). Oxaliplatin and SU-5416 was obtained from Sigma-Aldrich Chemical Co. (St Louis, MO, USA), EGFR kinase inhibitor Tyrphostin AG-1478 were purchased from Merck (Darmstadt, Germany), Gleevec (Novartis Pharmaceuticals, Basel, Switzerland) was kindly provided by P. Hohenberger (Mannheim, Germany). 5-Fluorouracil, recombinant active caspase-3 and all other chemicals were purchased from Sigma (St. Louis, MO) unless indicated otherwise.
Tumor cell culture maintenance
Wild-type C-26 murine colon-carcinoma cells (referred to as C-26 cells), syngenic to BALB/c mice, were provided by the American Type Culture Collection™ (Manassas VA). Cells were cultivated in DMEM medium supplemented with 10% (v/v) heat-inactivated fetal bovine serum, 100 U/ml penicillin, 100 μg/ml streptomycin and 1% (v/v) glutamine in a humidified atmosphere of 95% air and 5% CO2 at 37°C. Stock cultures were stored in liquid nitrogen and used for in vitro experiments within 5 passages. The C-26 cells transfected by tHcred-DEVD-EGFP (referred to as C-26-c3s cells) were maintained under the same conditions as the wild-type cells. For in vivo studies, cells were harvested from subconfluent cultures. Cell viability was determined by trypan blue exclusion. Cell number was adjusted to 1×106 cells in 0,5 ml PBS for intra-portal and intra-peritoneal injection to induce liver and peritoneal metastases in mice.
Transfection of C-26 cells with tHcred-DEVD-EGFP
C-26 cells were stably transfected with tHcred-DEVD-EGFP plasmid cDNA.29 Transfection of the construct into C-26 cells was carried out by using FuGENE 6 Transfection Reagent (Hoffmann-La Roche, Basel, Switzerland) according to the manufacturer's instructions. By limited dilution 6 clones were derived and expression levels were determined using FACS. The clone with the brightest EGFP fluorescence was used for further characterization.
Fluorescence lifetime imaging microscopy
Frequency domain fluorescence lifetime imaging microscopy (FLIM) was performed as previously described,29 where the fluorescence lifetimes (τ) were determined from the phase shift in the fluorescence emission. EGFP was excited with an argon 488 nm laser line modulated at a frequency of 80 MHz and the resulting fluorescence was separated from the excitation light using a combination of a dichronic beamsplitter (Q 490 LP; Chroma Technology Corp., Rockingham, VT) and a narrow band emission filter (HQ 510/20). The FLIM images were acquired on an Olympus IX 70 microscope (Olympus, Hauppauge, NY) equipped with a 60 X/1.4 NA oil immersion objective for observation of cultured cells. Fluorescence intensity images were captured using a scientific grade CCD camera (Photometrics Quantix, München, Germany). The data were analyzed using FLIM analysis software as previously described.30, 31 For FLIM procedures in mouse tissues see animal experiments.
In vitro experiments
C-26-c3s-cells were grown on 35mm glass bottom dishes and treated with 25 μM 5-FU for the indicated times. For microscopy, cells were washed and immersed in HEPES-buffered, phenol red- and riboflavin-free medium containing low carbonate (0.85 g/l). Afterwards live cell FLIM measurements were performed on cells adherent on the glass bottom of MaTek dishes.
For microinjection experiments, human recombinant caspase-3 was diluted in PBS at 0.1×10−3 U/ml, centrifuged (20 min, 15.000g, 4°C) and microinjected into the cytosol. Cells were microinjected using an Eppendorf microinjection system as previously described.32
Twelve-week-old female BALB/c mice were purchased from Janvier (France). They were bred under conditions of controlled temperature and light cycles with free access to food and water at the animal house of the Center for Medical Research (ZMF), University Heidelberg (University Clinic Mannheim). Experiments were performed in accordance with German legislation on the protection of animals and the guide, care, and use of laboratory animals (NIH Publication No. 86-23 revised edition, 1996).
Peritoneal metastases were generated by injecting C-26-c3s-murine colon carcinoma cells (1×106 cells in 0,5 ml PBS) into the peritoneal cavity. To produce liver metastasis the same number of cells in 100 μl PBS was injected into the portal vein. Here, mice were anaesthetized by i.p. injection of 16 mg/kg Xylazin (Rompun®) and 120 mg/kg Ketamin-Hydrochloride (Hostaket®) and a midline laparotomy was performed. Body temperature was maintained by a heat cabinet peri- and post-operatively. The portal vein was exposed and the cell suspension was slowly and carefully injected into the portal vein through 30-gauge needle. To ensure the postoperative fluid intake 1.5 ml saline solution was left in the abdominal cavity. The abdominal skin and musculature were closed in two layers with absorbable surgical sutures (Vicryl 5/0). To avoid possible pain stimuli in the early postoperative period 0,02 mg/kg Temgesic® (Buprenorphin) was injected subcutaneously just after closure of abdominal cavity. 32 tumor-bearing mice with peritoneal metastases and 32 mice with liver metastases were randomly divided in six groups (8 mice per group): (1) control (5 days of DMSO treatment); (2) 36 hours of 5-FU treatment; (3) 72 hours of 5-FU treatment; (4) 120 hours of 5-FU treatment (5) 180 hours of 5-FU treatment or (6) 72 hours of treatment and 48 hours no treatment. 5-FU was injected intraperitoneally ad 30 mg/kg body mass.33 By the end of the tenth day after tumor inoculation (consequently, after 36, 72 and 120 or 180 hours 5-FU treatment) animals were anesthetized as described above and midline laparotomy were performed through the scar tissue. Organs were mobilized, eventerated on glass cover slides and thus exposed for the in vivo FLIM experiments. The FLIM was performed as described above except that an 25×/0,8 water immersion objective was used to image cells on organ specimens. For determining the number of apoptotic cells by fluorescence lifetime imaging microscopy we used biopsy specimens taken from living mice. While the animals were kept under narcosis, about 10 pieces of tumor tissue were resected, put on the glass bottom of MaTek dishes and covered by a transparent cover slip. Since only small biopsy samples were taken only minor bleeding was observed. Care was taken to ensure that the exposed viscera were moistened adequately with warm saline solution throughout the experiments.
Re-isolation of cells from metastatic tissue
For re-isolation of the tumor cells from mice after 5 days of 5-FU and control mice, animals were anesthetized and laparotomized by the end of day 10 after i.p. tumor cell injection. The omental tumor tissue specimens were removed carefully from the abdominal cavity. Directly after removal, the tissue samples were divided into small parts using a scalpel on the culture flask. Tissue pieces were incubated in an atmosphere of 100% humidity and 5% CO2 at 37°C for 3 days in DMEM medium. The cells migrated away from tissue pieces onto the culture dishes. Thereafter, cells could be trypsinized and seeded onto MaTek dishes for FLIM experiments.
Quantitative Real Time PCR for EGFP
Quantitative Real Time PCR was performed on a LightCycler (Roche Molecular Biochemicals, Mannheim, Germany). Animals were sacrificed by cervical translocation under narcosis and the whole omentum or the whole liver was removed after midline laparotomy. Organs were homogenized and snap-frozen in liquid nitrogen and stored at −80°. DNA was isolated from tissue using the QIAamp DNA Mini tissue columns according to the manufacturer's instruction. The concentration and purity of the DNA samples were determined by spectrophotometry. 100 ng DNA from the omentum and 120 ng DNA from the liver was used for the PCR.
The reaction mixture consisted of 2 μl of LightCycler FastStart DNA Master SYBR Green I (FastStart Taq DNA polymerase, reaction buffer, dNTP mixture (with dUTP instead of dTTP), SYBR Green I dye and 10 mM MgCl2), 2.4 μl of 25 mM MgCl2, 0.5 mM of EGFP specific primers, 5′-TAC GGC AAG CTG ACC CTG AAG TTC-3′ (sense) and 5′-CGT CCT TGA AGA AGA TGG TGC-3′ (antisense)5, 36 and 2 μl of DNA derived from different preparations.
The cycling program consisted of 600 seconds initial denaturation at 95°C, 7 seconds 95°C denaturation, 64°C annealing for 5 seconds, and 72°C extension for 10 seconds with a transition rate of 20°C/second between temperature plateaus for a total of 35 cycles. Quantification data was analyzed using the LightCycler analysis software version 3.5. As standard the plasmid tHcred-DEVD-EGFP cDNA was used. The “Second Derivative Maximum Method” analysis algorithm was chosen for generating the standard curve. The error point was <0.1, the slope <3.3 Cp and the regression coefficient was r = −1.00. PCR products were analyzed on a 1% agarose gel to ensure specificity.
Data are displayed as box-plots showing means and standard deviation (s.d.). For the in vivo experiments means were determined for the measurements within one animal and these means were compared between the treatment groups. We compared each treatment group versus controls and other groups by one-way analysis of variance (ANOVA). If ANOVA indicated a significant difference between groups, pair wise multiple comparison of all means and post hoc testing using Tukey's method were employed to determine significant differences between groups and controls. Differences were considered significant if p < 0.05. SPSS 10.0.7 software (SPSS Inc. 1989–1999) was used for statistical analysis.
For in vivo-FRET imaging, we selected C-26 colorectal cancer cells (a highly malignant cell type in syngenic Balb/c mice) that were stably transfected with tHcred-DEVD-EGFP, a genetically encoded, optical FRET sensor for caspase-3 activity.29 These cells are further referred to as C-26-c3s-cells. We used fluorescence lifetime imaging microscopy (FLIM) as a robust method to quantify FRET signals from the sensor29 within living tumor tissues and specimens. Here, the fluorescence (excited-state) lifetime of the donor EGFP is shortened within the intact sensor construct due to the short range non-radiative transfer of energy to the acceptor tHcred. Upon cleavage of the caspase-3 sensor construct, this energy transfer cannot take place and EGFP reverts to its characteristic excited-state lifetime of ∼2.2ns. The functionality of the caspase-3 activity sensor was first tested by microinjection of active caspase-3 into C-26-c3s-cells. The non-injected cells exhibited a homogenously distributed fluorescence lifetime of EGFP in the sensor that was significantly shorter than the fluorescence lifetime of EGFP alone, demonstrating a clear FRET signal of the intact sensor throughout the cytoplasm. After microinjection of caspase-3, the cells showed an increase in fluorescence lifetime within minutes, close to that of free EGFP, which demonstrates the loss of the FRET signal and thereby complete sensor cleavage. C-26-c3s-cells were then treated with 5-FU in order to show that DNA-damage can induce apoptosis in these cells as measured by caspase-3 sensor cleavage. Under this 5-FU treatment in vitro, a clear population of apoptotic cells occurred after 48 h (p < 0.05, Fig. 1a) as indicated by their longer fluorescence lifetimes close to that of EGFP. Figure 1b shows representative images of untreated cells with low fluorescence lifetimes. Under treatment with 5-FU, apoptosis is induced and the fluorescence lifetimes increase.
We were now in the position to investigate chemotherapy induced apoptosis induction in vivo in these tumor cells. In vivo, interactions between the host microenvironment and the tumor graft determine tumor cell expression profiles,37 the level of growth factors and nutrients38 as well as angiogenesis and metastatic potential39 may all influence the ability of the cells to undergo apoptosis. We therefore created two syngenic murine tumor models. Figure 2a shows the experimental set up. Transfected cells were injected into the peritoneal cavity to simulate peritoneal carcinomatosis (Fig. 2b) or into the portal vein to simulate liver metastasis (Fig. 2c). After inoculation in mice, stably transfected cells showed parallel malignant attributes as compared to wild type cells. After introduction of 1×106 C-26-c3s-cells, the mice (n = 8) developed pronounced tumor spread in both models. Median survival of mice with fluorescent hepatic or peritoneal tumors were parallel to survival times observed for mice inoculated with wild type C-26 cells: 18.3 (±1.2) days for intraperitoneal versus 20 (±1.7) days for intrahepatic tumor spread.
To measure the apoptotic activity in untreated live tumor tissue 10 days after inoculation of C-26-c3s-cells, mice were laparotomized; the small bowel or the liver was eventerated onto a glass slide next to the living animal for FLIM. Fluorescent tumor cells could easily be detected on serosal surfaces using a 25×/0,8 water immersion objective. The measured fluorescence lifetimes corresponded to uncleaved caspase-3 sensor state. The homogenously distributed fluorescence lifetime values were stable over 60 min in the investigated living tissues. However, upon excising the tissues, the fluorescence lifetime values increased 15 min post mortem indicating the onset of apoptotic activity within the C-26-c3s-cells (Fig. 3). We therefore performed measurements in freshly resected tissues within the first 5 minutes. Importantly, no fluorescence could be detected in paraffin mounted fixed tissue serial slides derived from specimens of tumor bearing animals while tumor presence could be confirmed by hematoxylin-eosin staining. This indicated vulnerability of the fluorescent sensor to the fixation process.
We next analyzed in vivo response to chemotherapeutic apoptosis induction. Mice were treated for different times with 5-FU while the time of tumor presence within the animals was kept constant. Mice were treated with 5-FU ad 30 mg/kg i.p. over the indicated intervals, before animals were sacrificed. Fluorescence lifetimes determined in tissue samples from animals, which had obtained 5-FU treatment for 36 hours or 72 hours, increased to values above 2.0 ns (p < 0,05), indicating chemotherapy-induced apoptosis in human cells. Figure 4a shows the distributions of fluorescence lifetimes in the peritoneal metastasis group as a function of 5-FU treatment. Corresponding fluorescence lifetimes of the liver metastasis group are indicated in Figure 4d. Examplatory fluorescence lifetime images from which this data was derived are shown in Figure 4b. In parallel, post mortem, we could observe a decreased tumor load as determined by light cycler PCR for EGFP within the resected omentum (Fig. 4c; p < 0.05) or omentum (Fig. 4e; p < 0.05).
However, longer 5-FU treatment of mice led to fluorescence lifetimes in C-26-c3s-cells that were comparable to those in non-apoptotic tumor cells (Fig. 4a,d). Here, tumor loads were not different to shorter treatment intervals. This, as well as the lower proportion of apoptotic cells, indicated resistance to 5-FU induced apoptosis of the tumor cells in animals that were treated for longer than 72 h (Fig. 4c,e).
When treatment was applied for 72 hours and discontinued thereafter for the next 48 hours, no difference in tumor load could be observed as compared to animals, which were treated continuously over 120 hours or 180 hours (Fig. 4c). This data also indicates the selection of tumor cells that have become resistant to 5-FU induced apoptosis but have lost their proliferative attributes in the in vivo setting.
To further investigate the response of tumor cells which were exposed to 5-FU in vivo to secondary chemotherapy, we re-isolated tumor cells from treated and untreated tumor bearing animals. Tumor cells from mice, which were treated with 5-FU for 5 days, were re-isolated as primary culture. Upon incubation of these cells with 5-FU over 48 hours, no increase in fluorescence lifetimes and therefore no apoptosis could be observed (Fig. 5). The cells however showed an increase in fluorescence lifetime upon microinjected of active caspase-3, confirming that the sensor maintained its functionality after the in vivo passage (p < 0.05). This ex vivo experiment demonstrates that cells derived from treated animals did not have an apoptotic response upon reincubation with 5-FU. In contrast, cells that had been passaged through mice without 5-FU treatment displayed an ex vivo apoptotic response to 5-FU treatment comparable to cells before an in vivo passage (Fig. 5).
We have previously shown that in vitro cancer cell lines exhibit a high level of RTK phosphorylation40 and that RTK-inhibitors enhance the apoptotic effect of 5-FU.40 We therefore evaluated if growth factors might convey an anti-apoptotic survival signal. As expected, growth factor withdrawal in combination with 5-FU lead to apoptosis in the resistant cells (p < 0.05, Fig. 6a). Since many growth factors relay their signals via intracellular tyrosine kinase activities, we tested whether apoptosis could be induced in the resistant cell population by treatment with several clinically employed tyrosine kinase inhibitors in cancer treatment.41–43 In parallel, as a control, re-isolated cells from untreated animals were exposed to the compounds. While in the controls 5-FU in combination with other compounds induced apoptosis in 100% of the cells over 48 hours, we could show that only imatinib mesylate (Gleevec) could significantly increase the frequency of apoptosis in the resistant cells (p < 0.05). No significant increase in the frequency of apoptosis could be detected when comparing the EGFR inhibitors gefitinib (Iressa) or tyrphostin (AG1478) or the VEGFR inhibitor SU-5416 with 5-FU. Also treatment with oxaliplatin, another DNA-damaging agent which is frequently used clinically in 5-FU resistant tumors,44 did not lead to an increase in apoptosis (Fig. 6b).
The major problem in the treatment of cancer is not so much the eradication of the primary tumor, but rather the systemic spread by formation of surgically incurable metastases which are either primarily resistant to chemotherapy or develop resistance to therapy during treatment.45 In colorectal cancer, patient-mortality is particularly associated with the occurrence of metastases in the peritoneum and the liver.46, 47 Syngenic mouse models are powerful tools to simulate human colorectal cancer development and progression.5, 48 In these models, the effectiveness of chemotherapy can be assessed by the frequency of tumor cell apoptosis. Quantification of tumor cell apoptosis in tissues is presently based on histomorphologic and in-gel methods such as tunel, where sections for microscopy or specimens for homogenization are taken from tissue invasively, using biopsy or post mortem specimens.49–51
Until now, however, it has been difficult to directly observe molecular reactions in cells within organs of living laboratory animals such as the activation of proteases in apoptosis commitment. Here, we introduced an approach that permits the direct visualization and quantification of apoptotic activity in tumors of living animals. Employing FLIM, we could resolve chemotherapy-induced apoptosis in tumor dissemination to the liver and to the peritoneum as determined by caspase-3 activity.
We hereby show that, in principle, protease activities can be visualized in a small animal model. Especially for the observation of more dynamic processes imaging, quality could be further improved by the use of chamber models.57–58 Whole body FLIM-imaging could be applicable in future projects.59
Using FLIM, we could demonstrate that tHcred-DEVD-EGFP sensor-transfected C-26 cells undergo apoptosis when exposed to 5-FU in vitro. After inoculation in mice, we show that sensor transfected cells could be visualized on the serosal surfaces and apoptosis could be determined by reading out sensor state via fluorescence lifetimes. During 5-FU treatment in vivo, shortly after an initial treatment response, the development of apoptosis-resistant tumor cell subpopulations was observed by FLIM. Post mortem, the residual tumor load after treatment within the tissues could easily be determined using a light cycler PCR for EGFP cDNA as indicator of the tumor load. To investigate these cells further, cells were re-isolated from mice treated with 5-FU and from untreated controls. After re-isolation as primary culture, these cells remained unresponsive to 5-FU in vitro, in contrast to cells that were passaged through untreated mice. Here, we have evaluated apoptosis induction in these ex-vivo cells to evaluate secondary chemotherapy strategies. Interference with survival signalling is an appealing approach to the induction of apoptosis in tumor cells.29, 40 While the availability of growth factors is thought to define the size of various tissues by dictating the delicate balance between proliferation and apoptosis within a particular organ, limitations in growth factor availability and signalling in transformed cells leads to cell death.52 Therefore, growth factor withdrawal led to apoptosis when cells were re-exposed to 5-FU.
In order to mimic growth factor withdrawal in an environment that normally contains growth factors we used inhibitors of tyrosine kinases that normally relay growth factor signals inside the cell. While the tyrosine kinase inhibitors tested in combination with 5-FU induced apoptosis in 100% of the controls, only imatinib mesylate proved to be efficient in the re-induction of apoptosis in 5-FU resistant cells. Gleevec, a kinase inhibitor which has been developed to inhibit BCR-abl in chronic myeloid leukemia,53 is also an inhibitor of tyrosine kinases54 such as c-kit and PDGFR55 and exhibits a substantial therapeutic activity in patients with gastrointestinal stromal tumors,56 associated with constitutive activation of the c-kit tyrosine kinase. Recently, Gleevec was demonstrated to be effective in human and mouse colon cancer cells in vitro and in vivo.35
In summary, we show that FRET-based sensors can be successfully employed in a small-animal model to study syngenic cancer physiology and the development of chemotherapy resistance. The functional imaging techniques employed here can be useful to develop second line therapeutic approaches. Using a similar technical platform and sensors for other key activities implicated in cancer biology, these models could be transposed to other types of cancer to monitor drug therapies and allow the development of specific, disease adapted chemotherapy before and after development of metastasis, as well as for the case of resistance against the first line of treatment.
This work was supported by the EU grant “Insight Inside” (MK) and the Wilhelm Sander Stiftung (MK and VY). We thank H. Allgayer for helpful discussions.
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