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Research ArticleArticle

Lysosomal Sequestration Determines Intracellular Imatinib Levels

Herman Burger, Alexander T. den Dekker, Sandra Segeletz, Antonius W. M. Boersma, Peter de Bruijn, Maria Debiec-Rychter, Takahiro Taguchi, Stefan Sleijfer, Alex Sparreboom, Ron H. J. Mathijssen and Erik A. C. Wiemer
Molecular Pharmacology September 2015, 88 (3) 477-487; DOI: https://doi.org/10.1124/mol.114.097451

Abstract

The intracellular uptake and retention (IUR) of imatinib is reported to be controlled by the influx transporter SLC22A1 (organic cation transporter 1). We recently hypothesized that alternative uptake and/or retention mechanisms exist that determine intracellular imatinib levels. Here, we systematically investigate the nature of these mechanisms. Imatinib uptake in cells was quantitatively determined by liquid chromatography–tandem mass spectrometry. Fluorescent microscopy was used to establish subcellular localization of imatinib. Immunoblotting, cell cycle analyses, and apoptosis assays were performed to evaluate functional consequences of imatinib sequestration. Uptake experiments revealed high intracellular imatinib concentrations in HEK293, the leukemic cell lines K562 and SD-1, and a gastrointestinal stromal tumor cell line GIST-T1. We demonstrated that imatinib IUR is time-, dose-, temperature-, and energy-dependent and provide evidence that SLC22A1 and other potential imatinib transporters do not substantially contribute to the IUR of imatinib. Prazosin, amantadine, NH4Cl, and the vacuolar ATPase inhibitor bafilomycin A1 significantly decreased the IUR of imatinib and likely interfere with lysosomal retention and accumulation of imatinib. Costaining experiments with LysoTracker Red confirmed lysosomal sequestration of imatinib. Inhibition of the lysosomal sequestration had no effect on the inhibition of c-Kit signaling and imatinib-mediated cell cycle arrest but significantly increased apoptosis in imatinib-sensitive GIST-T1 cells. We conclude that intracellular imatinib levels are primarily determined by lysosomal sequestration and do not depend on SLC22A1 expression.

Introduction

Imatinib mesylate is successfully used in the treatment of chronic myeloid leukemia (CML) and gastrointestinal stromal tumors (GISTs). The intracellular uptake and retention (IUR) of imatinib in tumor cells is believed to be a critical factor for response, as it is considered to determine the effective imatinib concentration at the therapeutic targets within the tumor cell (Thomas et al., 2004; White et al., 2006; Wang et al., 2008). The IUR of imatinib relies on the delicate balance between drug uptake and efflux. Both influx and efflux transporters have been implicated in the active transport of imatinib into and out of cells (Eechoute et al., 2011). We have previously demonstrated that imatinib is a substrate of ABCB1 and ABCG2, which are efflux transporters modulating gastrointestinal absorption, distribution, and hepatic elimination of imatinib (Burger and Nooter, 2004; Burger et al., 2004, 2005). These efflux transporters are also expressed at the blood-brain barrier, where they inhibit the disposition of imatinib in the brain (Oostendorp et al., 2009). Whether ABCB1 and ABCG2 activity critically affects the IUR of imatinib in tumor cells and, as such, affects the outcome of imatinib treatment is still not clear (Mahon et al., 2000; Zong et al., 2005; Widmer et al., 2007).

The putative clinical significance of potential imatinib uptake transporters has extensively been investigated in the last decade as reflected by the numerous (pre-) clinical studies that have been conducted to investigate their role in the pharmacokinetics and/or pharmacodynamics of imatinib (Thomas et al., 2004; Crossman et al., 2005; White et al., 2006, 2007, 2010; Hu et al., 2008; Wang et al., 2008; White and Hughes, 2012; Nies et al., 2014). SLC22A1, also referred to as organic cation transporter (OCT) 1, has frequently been implicated in the intracellular uptake and disposition of imatinib in CML cells (Thomas et al., 2004; White et al., 2006), although controversy exists with regard to its precise role (Burger et al., 2013; Nies et al., 2014). SLC22A1 transporter expression/activity toward imatinib is reported to predict the long-term outcome of chronic-phase CML (White et al., 2006, 2007, 2010, 2012; Wang et al., 2008). Assessment of SLC22A1 transporter activity in cell lines and clinical CML samples was derived from imatinib IUR experiments in the presence or absence of SLC22A1 inhibitors such as prazosin or amantadine (Thomas et al., 2004; White et al., 2006). These functional SLC22A1 activity assays heavily depend on the specificity of these inhibitors. To definitely claim that SLC22A1 activity dictates the IUR of imatinib in CML cells, it is essential to confirm that these inhibitor-based assays truly reflect SLC22A1-associated transporter activity. This is a clinically relevant issue particularly since it has been proposed that SLC22A1 transporter activity should be used to appropriately identify patients with chronic-phase CML who are likely to respond poorly to imatinib (White and Hughes, 2012). We recently demonstrated that, irrespective of SLC22A1 levels, both prazosin and amantadine potently inhibit the IUR of imatinib (Burger et al., 2013). We therefore hypothesize that these compounds are not specific for SLC22A1 and interfere with alternative imatinib uptake/retention mechanisms. Here, we examined the IUR processes of imatinib in greater detail.

Materials and Methods

Chemicals and Reagents.

Prazosin, amantadine, carnitine, 1-methyl-4-phenylpyridinium (MPP+), sulforhodamine B, and ammonium chloride were obtained from Sigma-Aldrich (Zwijndrecht, The Netherlands). Other reagents and chemicals were purchased from the indicated suppliers: zosuquidar/LY335979 (Selleckchem, Houston, TX), elacridar/GF120918 (GlaxoSmithKline, Hertfordshire, UK), bafilomycin A1 (LC Laboratories, Woburn, MA), and 4-[4-(dimethylamino)styryl]-N-methylpyridinium iodide (ASP+; Molecular Probes, Eugene, OR). Imatinib mesylate was provided by Novartis Pharma AG (Basel, Switzerland).

Cell Lines and Culture Conditions.

All the cell lines used and their main characteristics are listed in the Supplemental Material (Supplemental Table 1). Human embryonic kidney (HEK) 293 cells transfected with the pcDNA3 vector alone (HEK293/Neo) or the pcDNA3/SLC22A1 gene (HEK293/SLC22A1) were provided by Dr. H. Koepsell (Institute of Anatomy and Cell Biology, Julius Maximilians University, Würzburg, Germany). All cell lines were cultured at 37°C and 5% CO2 in HEPES-buffered RPMI 1640 medium containing Glutamax (Gibco/Life Technologies, Bleiswijk, The Netherlands) supplemented with 10% (v/v) fetal bovine serum (Greiner, Alphen a/d Rijn, The Netherlands), 100 U/ml penicillin (Sigma-Aldrich), and 100 µg/ml streptomycin (Sigma-Aldrich).

Quantitative Real-Time Reverse-Transcription Polymerase Chain Reaction.

RNA isolation, cDNA synthesis, and real-time reverse-transcription polymerase chain reaction (RT-PCR) are detailed in the Supplemental Materials and Methods. Relative mRNA expression levels of nine different human transporter genes were measured in duplicate by real-time RT-PCR using Assay-On-Demand products (Applied Biosystems/Life Technologies; Supplemental Table 2).

Intracellular Imatinib Accumulation.

IUR studies of imatinib were performed at 37°C in serum-free uptake buffer containing 142 mM NaCl, 5 mM KCl, 1 mM K2HPO4, 1.2 mM MgSO4, 1.5 mM CaCl2, 5 mM glucose, and 12.5 mM HEPES (pH 7.3). Adherent cells were harvested by trypsinization, and nonadherent cells were taken directly from exponentially growing cell cultures. Each uptake experiment consisted of three independently exposed cell cultures that were separately processed for IUR measurements. In brief, cells were collected by centrifugation, washed twice with phosphate-buffered saline (PBS), and aliquots of 2 × 106 cells were resuspended in 1.5 ml of prewarmed uptake buffer. Cell suspensions were exposed to imatinib (1–100 µM) at 37°C for the indicated times, collected by centrifugation at 4°C, washed twice with PBS, and stored at −20°C until further processing. Imatinib concentrations in cell pellets and supernatants were determined by liquid chromatography–tandem mass spectrometry as described in the Supplemental Materials and Methods.

Flow Cytometry.

ASP+ accumulation was determined as described previously (Burger et al., 2010). In brief, 2 × 106 cells in 2 ml of serum-free RPMI 1640 without phenol red were incubated with ASP+ at 37°C for the indicated time periods in the presence or absence of a specific inhibitor/competitor. Fluorescence intensity of the cells was measured by flow cytometry (488-nm laser) and expressed in arbitrary units.

Fluorescence Microscopy.

Fluorescence imaging was performed on a Zeiss LSM-700 microscope (Carl Zeiss, Oberkochen, Germany) or a Leica TCS SP5 (Leica Microsystems, Eindhoven, The Netherlands). Adherent cells were grown on coverslips, washed with PBS, and incubated in 1 ml of uptake buffer supplemented with 30 µM imatinib alone or imatinib in combination with 60 nM LysoTracker Red DND-99 (Molecular Probes, Bleiswijk, The Netherlands). Cells growing in suspension were directly centrifuged on a coverslip at 700g for 10 minutes. Cells were typically preincubated with inhibitors/competitors for 30 minutes at 37°C prior to cotreatment with imatinib. Imatinib fluorescence was visualized using a DAPI band-pass filter (excitation: 405/20-nm wavelength; emission: 460/50 nm). LysoTracker Red DND-99 was visualized using an Alexa 555 band-pass filter (excitation: 546/12 nm; emission: 575–640 nm). Lasers for confocal microscopy were set to 405 nm for imatinib and 555 nm for LysoTracker Red DND-99. Confocal images (Z-stacks) were routinely analyzed by open source ImageJ software (http://imagej.nih.gov).

Protein Extraction and Immunoblotting.

GIST-T1 or K562 cells were preincubated for 30–60 minutes with prazosin (100 µM), NH4Cl (10 mM), or bafilomycin A1 (50 nM). Cell lysates, prepared in lysis buffer [50 mM Tris-HCl (pH 7.5), 50 mM NaCl, 10% glycerol, 1% NP40, 0.5% Na-deoxycholate, 1 mM Na3VO4, 20 mM NaF, 1 mM Pefabloc, and a cocktail of protease inhibitors], were cleared by centrifugation (20,000g) at 4°C for 15 minutes. Protein concentration was determined by the Bradford assay (BioRad, Veenendaal, The Netherlands). Equal amounts of protein (∼20 µg) were subjected to SDS-PAGE and transferred to a polyvinylidene fluoride membrane by electroblotting. Blots were blocked in Tris-buffered saline/Tween 20 (0.1% Tween 20) containing 5% nonfat dry milk. Primary antibodies, diluted in Tris-buffered saline/Tween 20 supplemented with 5% (w/v) bovine serum albumin (fraction V; Sigma-Aldrich), were as follows: rabbit monoclonal anti–CD117/c-Kit (YR145, 1:2500; Cell Marque, Rocklin, CA), rabbit polyclonal anti–phospho-c-Kit (Tyr719, 1:1000; Cell Signaling Technology, Leiden, The Netherlands), anti–phospho-Akt (Ser473, 1:2000; Cell Signaling Technology), anti–phospho-CrkL (Tyr207, 1:1000; Cell Signaling Technology), and mouse monoclonal anti-CrkL (32H4, 1:1000; Cell Signaling Technology). Mouse monoclonal anti–β-actin (AC-15, 1:5000; Sigma-Aldrich) was used as a loading control. Horseradish peroxidase–conjugated goat-anti-mouse (1:10,000; Santa Cruz Biotechnology, Heidelberg, Germany) or horseradish peroxidase–conjugated goat-anti-rabbit (1:10,000; Jackson Immunoresearch, Suffolk, UK) was used as a secondary antibody, and enhanced chemiluminescence (SuperSignal West Pico Chemiluminescent Substrate; Thermo Scientific, Rockford, IL) was used to visualize the signal on film.

Apoptosis Assay and Cell Cycle Analysis.

For the quantitation of apoptosis, untreated, imatinib-treated, NH4Cl-treated, and imatinib/NH4Cl-treated GIST-T1 cells were harvested and stained with fluorescein isothiocyanate–annexin V and propidium iodide (PI) using the fluorescein isothiocyanate–annexin V apoptosis detection kit I (BD Biosciences, Breda, The Netherlands), following the manufacturer’s protocol. For cell cycle analysis, untreated, imatinib-treated, NH4Cl-treated, and imatinib/NH4Cl-treated GIST-T1 cells were harvested and fixed in 70% ethanol for 15 minutes, washed in PBS, and resuspended in PBS/RNase/PI (20 μg/ml RNase, 50 μg/ml PI, 1% fetal bovine serum). Analysis was performed on a flow cytometer (FACScan; BD Biosciences) using a 488-nm laser with standard emission filters.

Statistical Analysis.

Accumulation data were analyzed and graphically expressed using GraphPad Prism version 5 (GraphPad Software, La Jolla, CA). Statistical significance was tested using an unpaired Student’s t test. Uptake data are expressed as the mean ± S.D. of at least three independently exposed cell cultures.

Results

Imatinib Interacts with SLC22A1.

We previously showed that ectopically expressed SLC22A1 (OCT1) in Xenopus laevis oocytes and HEK293 cells resulted in a consistent but limited increase in the IUR of imatinib (Hu et al., 2008; Burger et al., 2013). In the present study, we used the HEK293/Neo and HEK293/SLC22A1 cell lines (Burger et al., 2013) to further delineate the precise role of this uptake transporter in the IUR of imatinib. Flow cytometry showed that SLC22A1 overexpression in HEK293 cells resulted in a ∼10-fold enhanced intracellular concentration of ASP+, a fluorescent substrate of SLC22A1 (Fig. 1A). As the expression of SLC22A1 is low in HEK293/Neo cells, the uptake of ASP+ in HEK293/Neo is probably due to passive diffusion or SLC22A1-independent processes. Cotreatment with imatinib inhibited ASP+ uptake in a dose-dependent manner in these HEK293/SLC22A1 cells (Fig. 1A). The addition of 100 µM imatinib nearly blocked all of the SLC22A1-mediated uptake of ASP+.

Fig. 1.
Fig. 1.

Imatinib interacts with SLC22A1 (OCT1) and inhibits SLC22A1-mediated ASP+ uptake. (A, left panel) Analysis of ASP+ uptake in HEK293/SLC22A1 (shaded histogram) and HEK293/Neo control cells (open histogram). Cells were incubated with 1 µM ASP+ for 20 minutes at 37°C. Intracellular ASP+ accumulation was measured by FACScan flow cytometry. ASP+ levels were increased (∼10-fold) in the SLC22A1-overexpressing cells compared with the Neo control cells. (A, right panel) SLC22A1-mediated accumulation of ASP+ was inhibited in a dose-dependent manner. HEK293/SLC22A1 cells were coincubated at 37°C for 20 minutes with ASP+ (1 µM) and imatinib at the indicated concentrations diluted in serum-free RPMI 1640 without phenol red. The open (white) histogram represents the ASP+ accumulation of HEK293/Neo control cells. (B) Imatinib intracellular uptake and retention in HEK293/SLC22A1 and HEK293/Neo control cells. Uptake of imatinib (IM) was quantified by liquid chromatography–tandem mass spectrometry. Cells were exposed to 3 µM (upper panels) and 30 µM (lower panels) imatinib for 30 minutes (left panels) and 60 minutes (right panels) in prewarmed uptake buffer at 37°C. Imatinib uptake in HEK293/SLC22A1 cells was, on average, about 45% higher compared with HEK293/Neo control cells. Columns and error bars represent mean values ± S.D. of at least three independent IM-treated and separately processed cell cultures. Student’s t test was used to assess significant differences (*P < 0.05).

Next, we investigated whether imatinib is a substrate for SLC22A1, and as such, may function as a competitive inhibitor for other substrate drugs such as ASP+. Imatinib uptake experiments in which imatinib concentrations and exposure times were varied consistently indicated that the IUR of imatinib was only modestly increased (∼50%) in HEK293/SLC22A1 cells compared with HEK293/Neo control cells (Figs. 1B). Only in a single case was statistical significance (P < 0.05) reached. We conclude that, at least under these experimental conditions, imatinib is a rather weak substrate of SLC22A1 in addition to being an inhibitor of SLC22A1-mediated transport. It was observed that HEK293/Neo control cells, exhibiting relatively low SLC22A1 expression and activity compared with HEK293/SLC22A1 transfectants, already accumulate considerable levels of imatinib (Burger et al., 2013). To examine whether imatinib uptake in HEK293/Neo is mediated by constitutively expressed SLC22A1, we carried out an imatinib uptake experiment in the presence and absence of MPP+, an inhibitor of SLC22A1-3 (OCT1-3). MPP+ did not influence the IUR of imatinib (Supplemental Fig. 1), ruling out the involvement of SLC22A1 and indicating that other transporters, or alternative imatinib uptake or retention mechanisms, may play a role as well.

OCTN1/2 and ABC Transporter Expression in HEK293 Do Not Modulate the IUR of Imatinib.

The activity of other putative imatinib transporters might determine the IUR of imatinib in HEK293/Neo cells. Therefore, we analyzed the mRNA expression level of a number of candidate imatinib transporters (ABCB1, ABCG2, SLC22A1-5, SLCO1B1/3) by quantitative RT-PCR in these cells (Fig. 2A).

Fig. 2.
Fig. 2.

Transporter mRNA expression profiling. The relative mRNA expression of drug transporters implicated in imatinib transport was determined by real-time RT-PCR in HEK293/Neo (A), K562 (B), SD-1 (C), and GIST-T1 (D). mRNA expression levels were normalized to GAPDH and HPRT, and expressed in arbitrary units (A.U.). The normalized mRNA expression level of PBGD (first open bar), another well established housekeeper, is included for comparison. CT values were measured in duplicate, and those ≥35 were considered to be under the detection limit (<1 a.u.) and excluded. P-gp, P-glycoprotein.

The SLC22A4 (OCTN1) and SLC22A5 (OCTN2) solute carriers were expressed in HEK293/Neo cells (Fig. 2A). SLC22A5 transporter activity has previously been implicated in the IUR of imatinib (Hu et al., 2008). The related SLC22A4/5 are sodium-dependent transporters involved in the uptake of carnitine. We investigated whether carnitine interfered with the uptake of imatinib in HEK293/Neo cells. The presence of carnitine up to 100 µM had no significant effect on the IUR of imatinib in either HEK293/Neo or HEK293/SLC22A1 cells (Supplemental Fig. 2). This clearly suggests that, at least in HEK293 cells, OCTN activity does not contribute to the IUR of imatinib.

Both ABCB1 (MDR1) and ABCG2 (BCRP) are expressed in HEK293 cells (Fig. 2A). These membrane drug transporters are known to decrease the IUR of imatinib by active outward-directed transport (Burger et al., 2004, 2005). Zosuquidar is a selective inhibitor of P-glycoprotein (ABCB1), and elacridar is known to potently inhibit both P-glycoprotein and ABCG2 (Bihorel et al., 2007). However, preincubation with elacridar or zosuquidar did not increase intracellular imatinib levels in HEK293 cells (Supplemental Fig. 3). Apparently, these potential imatinib efflux transporters are not biologically active in HEK293, or their expression is too low to have a significant effect on the IUR of imatinib. We conclude that drug transporters for which evidence exists in the literature to functionally link them to imatinib transport are not operational in HEK293 cells.

Quantitative Measurements Reveal High Intracellular Imatinib Concentrations.

A quantitative evaluation of the HEK293 imatinib IUR experiments revealed that a substantial part (17–20%) of the administered imatinib was actually recovered in the cellular fraction (Supplemental Table 3). Apparently, imatinib is rapidly taken up in the cells, where it is retained and accumulates. Based on a total cell volume of 5 μl for 2 × 106 cells (le Coutre et al., 2004) and a total incubation volume of 1.5 ml, we could estimate the intracellular imatinib concentration to be ∼70-fold higher, approaching the millimolar range, than the imatinib concentration in the incubation medium (Supplemental Table 3). Similar observations were made in other cell lines: intracellular imatinib concentrations were found to be 35- to >100-fold higher than those in the incubation medium. Our findings support reports in the literature that describe high intracellular imatinib levels in peripheral blood mononuclear cells from CML patients (Bouchet et al., 2013), in leukemic cell lines K562 and KCL22 (le Coutre et al., 2004), and in HL-60 cells (Bourgne et al., 2012). To explain these findings, most studies infer the existence of an active uptake mechanism involving a saturable drug transporter (Bourgne et al., 2012; Bouchet et al., 2013). Alternatively, one can envisage the existence of a highly efficient intracellular mechanism capable of accumulating and retaining imatinib.

IUR of Imatinib Is Time-, Dose-, Temperature-, and Energy-Dependent.

To obtain more insight in the cellular uptake process of imatinib, we investigated the uptake kinetics and dose, temperature, and energy dependency. In addition to HEK293 cells, we included two leukemic cell lines, i.e., K562 expressing a p210 BCR-ABL transcript typical for CML, and the SD-1 cell line expressing a p190 BCR-ABL transcript typical for acute lymphoblastic leukemia. Both cell lines are derived from leukemias that are routinely treated with imatinib similar to GISTs, the tumor type from which the GIST-T1 cell line has been established. The potential imatinib transporter profile of K562 and SD-1 is similar to that of HEK293, with the exception that K562 cells express low levels of SLCO1B1 (Fig. 2).

We determined the IUR of imatinib in HEK293/Neo (Fig. 3A) and K562 (Fig. 3B) as a function of time. Intracellular accumulation of imatinib is time-dependent, displaying an initial rapid uptake within the first 15 minutes, which leveled off thereafter. Similar imatinib uptake kinetics were shown for HEK293/SLC22A1 (Supplemental Fig. 4). Next, we showed that the intracellular accumulation of imatinib is dose-dependent in HEK293/Neo (Fig. 3C), K562 (Fig. 3D), and SD-1 cells (Fig. 3E), which suggests a saturable mechanism.

Fig. 3.
Fig. 3.

The intracellular uptake and retention of imatinib is time-, dose-, and energy-dependent. Time-dependent uptake of imatinib in HEK293/Neo (A) and K562 cells (B). Cells were exposed to 10 µM imatinib for the indicated time periods in uptake buffer at 37°C. Uptake of imatinib (IM) was quantified by liquid chromatography–tandem mass spectrometry. Concentration-dependent uptake of imatinib in HEK293/Neo (C), K562 (D), and SD-1 (E) cells. Cells were exposed to the indicated imatinib concentrations for 30 minutes. (F) Imatinib uptake in HEK293/Neo cells is temperature-dependent. Cells were exposed to imatinib (10 µM) in uptake buffer and incubated at the indicated temperatures (37°C versus 4°C) for 30 minutes. (G) The IUR of imatinib is an active energy (ATP)–dependent process. ATP depletion was established by sequential depletion of glucose and subsequent addition of NaN3. HEK293/Neo cells were preincubated for 1 hour in prewarmed uptake buffer (control), glucose-free uptake buffer, or glucose-free uptake buffer containing NaN3 (10 mM) prior to imatinib (10 µM) exposure. Columns and error bars represent mean values ± S.D. of at least three independent IM-treated and separately processed cell cultures. Student’s t test was used to assess significant differences. **P < 0.01; ***P < 0.001.

To investigate whether the IUR of imatinib involves an active energy-dependent process, we compared the intracellular imatinib levels in HEK293/Neo cells at 4 and 37°C. The IUR of imatinib in cells exposed to 10 µM imatinib at 4°C was about 27-fold lower than that observed at 37°C (Fig. 3F), pointing toward an active uptake/retention mechanism instead of passive diffusion. Next, we determined the effect of energy depletion by inhibiting ATP production through glycolysis and mitochondrial oxidative phosphorylation. Depletion of glucose in HEK293/Neo cells treated with 10 µM imatinib resulted in a more than 2-fold decrease in IUR, and the sequential addition of NaN3, an inhibitor of oxidative phosphorylation, led to a total 9-fold decrease (Fig. 3G). These results clearly demonstrate that the IUR of imatinib is an active and energy-dependent process.

Lysosomotropic Compounds Decrease the IUR of Imatinib.

To elucidate the mechanisms that drive the IUR of imatinib, we focused on known inhibitors of the imatinib uptake process. Prazosin and amantadine significantly decreased the IUR of imatinib not only in HEK293/Neo and HEK293/SLC22A1 cells, but also in the leukemic K562 (Burger et al., 2013), SD-1 cells (Supplemental Fig. 5), and notably in GIST-T1 cells (Fig. 4C) that do not express SLC22A1 (Fig. 2D). Clearly, the effects of prazosin and amantadine on the IUR of imatinib could not be explained by inhibition of SLC22A1-mediated transport of imatinib (Burger et al., 2013).

Fig. 4.
Fig. 4.

Bafilomycin A1 and NH4Cl reduce the intracellular uptake and retention of imatinib. The effect of bafilomycin A1 (A and C) and NH4Cl (B and C) on the IUR of imatinib in HEK293/Neo, K562, SD-1, and GIST-T1. Cells were incubated for 1 hour with the indicated concentrations of bafilomycin A1 and the lysosomotropic NH4Cl and thereupon treated for an additional 30 minutes with 10 µM imatinib, with the exception of the cells in (B) (HEK293/Neo, top panel) and (C), which were treated with 3 and 1 µM imatinib, respectively. (C) Effect of bafilomycin A1 (50 nM), prazosin (100 µM), and NH4Cl (10 mM) on the intracellular accumulation of imatinib in GIST-T1 cells. Uptake of imatinib (IM) was determined by liquid chromatography–tandem mass spectrometry. Columns and error bars represent mean values ± S.D. of at least three independent imatinib-treated and separately processed cell cultures. Student’s t test was used to assess significant differences. *P < 0.05; **P < 0.01; ***P < 0.001.

As amantadine accumulates preferentially in lysosomes (Kornhuber et al., 2010), we reasoned that amantadine may prevent imatinib from being sequestered into lysosomes, thereby decreasing the IUR of imatinib. Therefore, we investigated whether the IUR of imatinib in HEK293/Neo, K562, SD-1, and GIST-T1 cells was inhibited by bafilomycin A1, a strong inhibitor of the vacuolar type H(+)-ATPase (Mattsson et al., 1991), and NH4Cl, a well known lysosomotropic compound. Bafilomycin A1 significantly decreased the IUR of imatinib in a dose-dependent manner by preventing the acidification of the lysosomes (Fig. 4A). NH4Cl, a weak base that rapidly increases lysosomal pH, also decreased the IUR of imatinib (Fig. 4B). Finally, we showed that bafilomycin A1 and NH4Cl significantly decreased imatinib uptake in GIST-T1, an imatinib-sensitive GIST cell line expressing a mutated exon 11 c-Kit variant (Fig. 4C). These results strongly suggest that imatinib is a lysosomotropic agent itself accumulating in acidic lysosomal compartments.

Imatinib Is Effectively Sequestered in Lysosomes.

To visualize cellular imatinib uptake and prove that imatinib is sequestered in lysosomes, we exploited the fluorescent properties of imatinib. Imatinib is a weak fluorescent compound with maximal absorption and emission at ∼290 and ∼540 nm, respectively, and displays a pH-independent linear relationship between concentration in the millimolar range and fluorescence (Supplemental Fig. 6). Fluorescence microscopy revealed that imatinib fluorescence is not homogenously distributed within the cytoplasm but displayed a clear punctuated fluorescent pattern predominantly in the perinuclear region of imatinib-treated HEK293/Neo and lung-derived H226 cells (Fig. 5A; Supplemental Fig. 7). A similar fluorescent pattern, suggesting vesicular accumulation of imatinib, was also observed in leukemic cell lines as well as ovarian and breast cancer cell lines (Fig. 5B).

Fig. 5.
Fig. 5.

The fluorescent detection of imatinib in the lysosomal compartment. (A, left panel) Confocal fluorescence imaging of imatinib in HEK293/Neo cells incubated with 30 µM imatinib for 30 minutes at 37°C. Depicted are a bright field image and imatinib fluorescence in green (left panel). (A, right panel) Fluorescence imaging of imatinib (green), LysoTracker Red DND-99 (red), and overlay (yellow) in H226 cells coincubated with 30 µM imatinib and 60 nM LysoTracker Red for 30 minutes at 37°C. The LysoTracker Red fluorescence shows a complete overlap with the imatinib fluorescence signal, indicating that imatinib is sequestered in lysosomes. (B) Confocal fluorescence microscopy showing lysosomal sequestration of imatinib in various cancer cell lines. Cells were incubated with 30 µM imatinib and 60 nM LysoTracker Red for 30 minutes at 37°C. Depicted are fluorescent images of imatinib (green signal, top panels), LysoTracker Red DND-99 (red signal, middle panels), as well as the merged images (bottom panels). Scale bar: 15 µm. (C) Effect of bafilomycin A1, prazosin, and NH4Cl on the lysosomal sequestration of imatinib in HEK293/Neo cells. Lysosomal sequestration of imatinib (left panel, green signal) was completely inhibited, as judged by the absence of a fluorescent signal, by the addition of 50 nM bafilomycin A1, 100 µM prazosin, and 10 mM NH4Cl (respective panels to the left). Cells were preincubated for 1 hour at 37°C with the respective compounds prior to imatinib treatment (30 µM; 30 minutes at 37°C). Scale bar: 10 µm.

To demonstrate that the weak base imatinib is sequestered in acidic lysosomes, we coincubated H226 cells with imatinib and LysoTracker Red DND-99, a viable fluorophore readily entering acidic subcellular organelles such as lysosomes (Nadanaciva et al., 2011). Confocal imaging clearly showed that the red fluorescent pattern from LysoTracker Red colocalized with the imatinib-associated green fluorescent signal as indicated by the yellow color in the overlay (Fig. 5A). Imatinib and LysoTracker Red stained the same subcellular structures in all cell lines examined, confirming that imatinib indeed accumulates in lysosomes (Fig. 5, A and B).

Time-lapse experiments to investigate the kinetics of lysosomal sequestration of imatinib showed that the uptake into the lysosomal compartment occurs rapidly, as the particulate imatinib fluorescence was observed within minutes (Supplemental Fig. 8A). Furthermore, a pulse-chase experiment was performed showing that the imatinib fluorescence remains visible for at least 12 hours (Supplemental Fig. 8B). IUR data measured by liquid chromatography–tandem mass spectrometry confirmed that imatinib is retained intracellularly, at least within the first hour after a washout (Supplemental Fig. 9A), and afterward declined to ∼70% at t = 2 hours and 30% at t = 8 hours (Supplemental Fig. 9B), as judged by fluorescence microscopy. Notably, bafilomycin A1, NH4Cl, and prazosin completely blocked lysosomal sequestration of imatinib (Fig. 5C). Altogether, we clearly showed that imatinib readily accumulates in lysosomes irrespective of cell origin and/or tumor type.

Effect of Lysosomal Sequestration on the Functional Activity of Imatinib.

The IUR of imatinib is reported to be an important factor for response, as it is believed to determine the effective imatinib concentration at the therapeutic target site (Thomas et al., 2004; White et al., 2006, 2010; Wang et al., 2008; Bouchet et al., 2013). A possible consequence of the rapid and efficient lysosomal sequestration of imatinib, which primarily determines the IUR of imatinib, could be that most of the imatinib in cells will not be able to adequately reach and interact with its molecular targets localized in other cell compartments. To examine this, we determined the functional activity of imatinib after modulating the lysosomal sequestration of imatinib. Imatinib-sensitive GIST-T1 cells express a mutant c-Kit encoding a constitutively activated c-Kit oncoprotein. Cotreatment of GIST-T1 with imatinib and prazosin (100 µM), bafilomycin A1 (50 nM), and NH4Cl (10 mM) effectively inhibited the phosphorylation of c-Kit and its downstream effector Akt (Fig. 6A). It should be noted that the respective concentrations of these compounds dramatically decreased (∼9-fold) the IUR of imatinib in these GIST-T1 cells (Fig. 4C). Notably, prazosin, bafilomycin A1, and NH4Cl on their own had no effect on the phosphorylation status of c-Kit and Akt (Fig. 6A). Treatment of GIST-T1 cells with imatinib (1 µM) only also clearly inhibited c-Kit signaling (Fig. 6A). Our findings indicate that, despite lysosomal sequestration, the imatinib levels within the GIST-T1 cells are adequate to effectively inhibit c-Kit in GIST-T1 cells. Similar results were obtained when the lysosomal sequestration of imatinib was modulated in K562 cells and the inhibition of BCR-ABL signaling was monitored by the phospho-CrkL levels (Supplemental Fig. 10).

Fig. 6.
Fig. 6.

Inhibition of lysosomal sequestration of imatinib and its effects on c-Kit inhibition, apoptosis, and cell cycle progression. (A) Western blot analysis of key signaling proteins in imatinib-sensitive GIST-T1 cells; c-Kit protein levels, phospho–c-Kit levels, and downstream effector Akt (phospho-Akt) levels were determined in the presence (+) and absence (−) of the indicated compounds. GIST-T1 cells were preincubated for 1 hour with prazosin (100 µM), bafilomycin A1 (50 nM), or NH4Cl (10 mM) and subsequently treated for an additional 6 hours with 1 µM imatinib. Equal amounts of protein were loaded, and β-actin was used as a loading control. (B) Effect of NH4Cl on imatinib-induced apoptosis in GIST-T1 cells. Apoptosis was quantified by flow cytometry using annexin V/PI stainings (Supplemental Fig. 11). Viable cells are represented by the annexin V−/PI− fraction (purple), early apoptotic cells by the annexin V+/PI− fraction (green), late apoptotic cells by the annexin V+/PI+ fraction (red), and dead cells by the annexin V−/PI+ fraction (blue). Cumulative percentages are shown. (C) Effect of NH4Cl on imatinib-induced cell cycle arrest in GIST-T1 cells. Cell cycle profiles were determined in untreated, NH4Cl-treated, imatinib-treated, and imatinib/NH4Cl cotreated GIST-T1 cells by FACS analysis (Supplemental Fig. 11). Cumulative percentages of sub-G1 (blue), G1 (red), S-phase (green), and G2/M (purple) are shown.

Next, we investigated whether conditions that prevent lysosomal accumulation of imatinib affected the cell cycle arrest observed in GIST-T1 cells and promoted apoptosis (Gupta et al., 2010). We noted that GIST-T1 cells could not endure prolonged exposure (48 hours) to bafilomycin A1 and NH4Cl; therefore, GIST-T1 cells were exposed for 2 hours to 2.5 μM imatinib, 10 mM NH4Cl, or a combination of both compounds, after which they were allowed to recover for 48 hours. Subsequently, we quantified the percentage of apoptotic cells in each of the conditions (Fig. 6B; Supplemental Fig. 11). As expected, imatinib induced apoptosis, whereas NH4Cl on its own did not promote the induction of apoptosis. Cotreatment of the cells with imatinib and NH4Cl resulted in the highest apoptotic levels, indicating a synergistic interaction between these compounds. We also examined the cell cycle profile in GIST-T1 cells under these conditions, observing that imatinib treatment alone induced a G1 arrest that did not significantly differ in the presence of NH4Cl (Fig. 6C; Supplemental Fig. 11). Our findings indicate that there is no apparent change in imatinib-induced G1 arrest and the inhibition of c-Kit signaling whether imatinib is present or absent in the lysosomes. In contrast, we clearly established that a reduced lysosomal sequestration of imatinib renders GIST-T1 cells more susceptible to apoptosis induction.

Discussion

SLC22A1 has been suggested to largely determine the IUR of imatinib, and its expression/activity are thought to predict the long-term outcome of chronic-phase CML cells (Thomas et al., 2004; Crossman et al., 2005; White et al., 2006, 2007, 2010; Wang et al., 2008; White and Hughes, 2012). SLC22A1 transporter activity is commonly derived from the IUR of imatinib with and without “specific” SLC22A1 inhibitors such as prazosin or amantadine. However, we showed that these inhibitors do not exclusively inhibit the SLC22A1 transporter (Burger et al., 2013). Here, we demonstrate that imatinib is sequestered in the lysosomal compartment, and that this process can be inhibited by prazosin and amantadine. We further conclude that intracellular imatinib levels are primarily determined by lysosomal sequestration and do not depend on SLC22A1 expression.

The present study shows that SLC22A1 has some activity toward imatinib, but does not significantly contribute to the IUR of imatinib in cancer cells. In addition to being a weak SLC22A1 substrate, imatinib clearly inhibits the SLC22A1-mediated transport of ASP+ and metformin (Minematsu and Giacomini, 2011). Apart from SLC22A1, other uptake transporters, including SLCO1B3, SLC22A4 (OCTN1), and SLC22A5 (OCTN2), have been implicated in the uptake and disposition of imatinib (Hu et al., 2008; Eechoute et al., 2011; Yamakawa et al., 2011). Polymorphisms in OCTNs were found to be associated with clinical response parameters in imatinib-treated GIST and CML patients (Angelini et al., 2013a,b). However, there is no compelling evidence that OCTNs are actively involved in imatinib transport, corroborating our own findings in HEK293 cells that carnitine, a high-affinity OCTN substrate, does not interfere with the IUR of imatinib. Furthermore, overexpression of SLC22A4 in COS-7 cells had no effect on the IUR of imatinib (McWeeney et al., 2010). Altogether, these data indicate that none of the known imatinib transporters plays a prominent role in the IUR of imatinib in HEK293 cells. Instead, our findings demonstrate that the IUR of imatinib is primarily determined by lysosomal sequestration. Accumulation of imatinib in lysosomes was described before by Chapuy et al. (2009) and, more recently, by Fu et al. (2014).

How can the reported association between “prazosin-defined OCT1 activity” and imatinib response in CML be explained? First, it is noted that the concentration of prazosin, or amantadine, used in these studies was not high enough to completely inhibit lysosomal sequestration of imatinib. The resulting variable levels of intracellular/lysosomal imatinib may have significantly confounded the interpretation. Second, prazosin—in addition to being a lysosomotropic agent—affects the cell in other ways as well; this is exemplified by Zhang et al. (2012), who show that prazosin rapidly perturbs endocytic dynamics. We also observed that prolonged exposure of K562 cells to prazosin alone already interferes with BCR-ABL signaling as judged by the reduced phospho-CrkL levels (Supplemental Fig. 10).

The physicochemical properties of imatinib (hydrophobic log P = 4.5 and pKa = 8.0) indicate that it is a weak base and rather lipid soluble. Weak basic compounds are generally membrane-permeable in their neutral form and therefore readily diffuse across biologic membranes. The parallel artificial membrane permeability assay, predicting the extent of passive transcellular permeability of a drug, indeed showed that imatinib easily crosses a synthetic lipid bilayer (Zimmerman et al., 2013). The pH difference between the cytosol (pH ∼7.2) and the lysosomal compartment (pH ∼5) is the driving force for lysosomal drug sequestration, and is maintained by membrane-bound ATP-dependent lysosomal proton pumps of the highly conserved vacuolar ATPase (V-ATPase) family (Forgac, 2007). Interestingly, loss of V-ATPase activity by gene disruption in yeast affected the tolerance for imatinib, providing evidence for an intimate link between imatinib and the vacuole in yeast, the yeast ortholog of the mammalian lysosome (dos Santos and Sa-Correia, 2009). The accumulation of imatinib in the lysosome may raise the lysosomal pH and thereby decrease the driving force for imatinib uptake into the lysosome, explaining the apparent saturability of the process (Fig. 3, C–E). Sequestration of imatinib in acidic lysosomes requires the passage over both the plasma and lysosomal membrane. It has been suggested that drug transfer directly from the cytosol into the luminal space of lysosomes may proceed via three different pathways: passive diffusion, autophagocytosis, and/or transporter-mediated accumulation (Kaufmann and Krise, 2007). Since passive diffusion has the highest accumulation capacity and is most efficient, we and others (Fu et al., 2014) speculate that the uptake into lysosomes likely proceeds by simple diffusion, although other mechanisms cannot be excluded. Within the lysosome, imatinib will be effectively protonated and become membrane-impermeable (Szakacs et al., 2005). This process, called ion trapping, requires the continuous acidification of the lysosomes by V-ATPase activity, explaining why the IUR of imatinib was found to be energy- and temperature-dependent. Furthermore, the definite amount of imatinib in lysosomes is likely confined, accounting for the observation that the IUR of imatinib is saturable. In line with our observations, it has been reported that drugs that sequester to lysosomes can reach final intracellular concentrations that are much higher than the drug concentrations in the surrounding medium (Kornhuber et al., 2010; Fu et al., 2014).

The physicochemical properties of amantadine and prazosin predict that they are potential lysosomotropic compounds that may function as a competitive inhibitor interfering with the lysosomal sequestration of imatinib. One may predict that the concomitant administration of imatinib and prazosin or other lysosomotropic drugs may significantly alter the pharmacokinetics and/or pharmacodynamics of the drugs.

How does the accumulation of imatinib in the lysosomal compartment interfere with its anticancer activity? We noted in GIST-T1 cells that c-Kit signaling is still inhibited, irrespective of whether imatinib is sequestered in lysosomes or not. However, when we focused on imatinib-induced cell cycle arrest and apoptosis, we observed that cotreatment of imatinib and lysosomotropic NH4Cl resulted in increased apoptosis. This phenomenon has been reported before in GIST (Gupta et al., 2010) and CML (Bellodi et al., 2009). The explanation may be that imatinib induces autophagy (Ertmer et al., 2007) as a survival mechanism. Autophagy is inhibited by the addition of strong lysosomotropic agents such as chloroquine or NH4Cl, causing increased cell death. There are indications that high intracellular levels of imatinib, or other tyrosine kinase inhibitors, entrapped in lysosomes for prolonged periods of time may lead to lysosomal dysfunction, including lysosomal membrane permeabilization, impaired autophagy, and eventually lysosomal-mediated cell death (Ellegaard et al., 2013; Groth-Pedersen and Jaattela, 2013). It is conceivable that the biologic activity and anticancer efficacy of imatinib are in part mediated through the lysosomal compartment. It has been suggested that lysosomal targeting may be a promising novel strategy to eradicate imatinib-resistant CML (Puissant et al., 2010). In addition, the accumulation of imatinib in lysosomes of healthy cells may underlie several side effects. For example, imatinib has been associated with congestive heart failure: although the incidence in clinical trials was low and typically occurred in patients with a pre-existing history of heart disease, patients with severe congestive heart failure have been reported while receiving imatinib (Kerkela et al., 2006). Imatinib-induced cardiac toxicity, observed in rodents and mechanistically linked to the physicochemical properties of imatinib, was described to be caused by efficient lysosomal sequestration of imatinib leading to lysosomal dysfunction and autophagy perturbation in cardiomyocytes (Herman et al., 2011; Hu et al., 2012). It may well be that other rare, or more frequently encountered, imatinib-related side effects are a direct consequence of a lysosomal impairment caused by the lysosomal accumulation of imatinib in healthy cells. This is a relevant clinical issue considering the prolonged daily use of imatinib by a large group of cancer patients. Moreover, treatment efficacy and/or treatment-related toxicity may be seriously affected by the fact that cancer patients frequently use multiple drugs simultaneously, a number of which may interfere with the lysosomal sequestration of imatinib.

It is clear that the lysosomal sequestration of imatinib as the main determinant of intracellular imatinib levels has been largely overlooked by the mainstream literature on this topic (Thomas et al., 2004; Crossman et al., 2005; White et al., 2006, 2007, 2010; White and Hughes, 2012). Instead, these reports mainly focused on the putative transporter-mediated uptake of imatinib notably by SLC22A1, ignoring the physicochemical properties of imatinib and the rather poor specificity of the inhibitors used to determine SLC22A1 activity. Although several papers describe a correlation between intracellular imatinib levels and treatment outcome (White et al., 2006; Bouchet et al., 2013) or between SLC22A1 activity, measured using prazosin and amantadine, and response (White et al., 2007, 2010), it may well be that, in reality, they have uncovered a correlation between the activity of the lysosomal compartment and treatment outcome. Recently, additional evidence was presented that the cellular uptake of imatinib into leukemic cells is independent of SLC22A1 (Nies et al., 2014), corroborating our findings. The authors hypothesized that other not yet identified transporters may be involved. We, however, identified lysosomal sequestration to be the main determinant for cellular imatinib uptake.

Acknowledgments

The authors thank Dr. H. Koepsell (Institute of Anatomy and Cell Biology, Julius Maximilians University, Würzburg, Germany) for providing the HEK293/SLC22A1-3 and HEK293/Neo control cells. The authors thank Solmaz Karakaya for technical assistance, and Tsion Abraham and Alex Nigg are acknowledged for their assistance with fluorescence (confocal) microscopy.

Authorship Contributions

Participated in research design: Burger, Wiemer.

Conducted experiments: Burger, den Dekker, Segeletz, Boersma, de Bruijn, Wiemer.

Contributed new reagents or analytic tools: Burger, den Dekker, Segeletz, Boersma, de Bruijn, Wiemer.

Performed data analysis: Burger, den Dekker, Segeletz, Boersma, de Bruijn, Wiemer.

Wrote or contributed to the writing of the manuscript: Burger, den Dekker, Segeletz, Boersma, de Bruijn, Debiec-Rychter, Taguchi, Sleijfer, Sparreboom, Mathijssen, Wiemer.

Footnotes

Abbreviations

ASP+
4-[4-(dimethylamino)styryl]-N-methylpyridinium iodide
CML
chronic myeloid leukemia
GIST
gastrointestinal stromal tumor
HEK
human embryonic kidney
IUR
intracellular uptake and retention
MPP+
1-methyl-4-phenylpyridinium
OCT
organic cation transporter
PBS
phosphate-buffered saline
PI
propidium iodide
RT-PCR
reverse-transcription polymerase chain reaction
V-ATPase
vacuolar ATPase

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Lysosomal Sequestration of Imatinib

Herman Burger, Alexander T. den Dekker, Sandra Segeletz, Antonius W. M. Boersma, Peter de Bruijn, Maria Debiec-Rychter, Takahiro Taguchi, Stefan Sleijfer, Alex Sparreboom, Ron H. J. Mathijssen and Erik A. C. Wiemer
Molecular Pharmacology September 1, 2015, 88 (3) 477-487; DOI: https://doi.org/10.1124/mol.114.097451
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