Adipocytes Sequester and Metabolize the Chemotherapeutic Daunorubicin Free

It is well established that obesity increases the risk for cancer mortality (1). While obesity has been linked with poorer outcome from several cancers, including that of the breast (2), colon (3), ovary (4), and prostate (5), no mechanisms have been proven to explain these effects. One potential contributor to poor cancer outcome in obesity could be the inadequacy of chemotherapy dosing. Excess adiposity can lead to alterations in chemotherapy pharmacokinetics. Lipophilic chemotherapies can preferentially accumulate in adipose tissue (6), thus increasing the volume of distribution, and reducing cancer cells exposure to the chemotherapy. Practices of dose capping, dosing by body surface area, and adjusting for ideal or lean body weight could further contribute to underdosing in obese patients. However, few studies have systematically evaluated how obesity alters the disposition of chemotherapies in patients.

Anthracyclines such as daunorubicin and doxorubicin are important chemotherapy agents used in a wide variety of cancers in children and adults. We have shown that adipocytes protect acute lymphoblastic leukemia (ALL) cells from a variety of chemotherapies, including daunorubicin (7, 8). We report herein that adipocytes sequester daunorubicin and metabolize it to an inactive form, showing for the first time that adipose tissue is a drug-metabolizing organ.

Daunorubicin and doxorubicin were purchased from Sigma Chemical Company. Daunorubicinol and doxorubicinol were purchased from Santa Cruz Biotechnology. Media and supplements were obtained from Gibco, Thermo Fisher Scientific. FBS was purchased from Denville Scientific.

Human ALL cell lines BV173 (Ph⁺ pre-B-ALL) and SEM (t4;11 pre-B-ALL) were acquired from ATCC, authenticated by short tandem repeats by the University of Arizona Genetics core in November 2016, and tested negative for mycoplasma. ALL cell lines were cultured in RPMI1640 containing 10% FBS, 1% sodium pyruvate, 1% Glutamax, and 0.1% gentamicin (“complete media”), and maintained at densities between 0.5 and 2.0 × 10⁶ cells per mL in a humidified incubator with 5% CO₂. The murine preadipocyte cell line 3T3-L1 from ATCC was differentiated into adipocytes as described previously (9) and used for experiments between days +7 and +11 of differentiation. As a control, undifferentiated 3T3-L1 cells were irradiated with 90 Gy to induce senescence and plated at confluence, referred herein as 3T3-L1 fibroblasts. Immortalized human adipocytes (ChubS7) were also differentiated and cultured as described previously. Human bone marrow–derived mesenchymal stem cells (MSC) were obtained from Thermo Fisher Scientific and differentiated as per manufacturer's instructions with MesenPRO medium. In coculture experiments, approximately 2 × 10⁵ ALL cells were cultured in 0.4-μm pore-size polycarbonate Transwell (Corning, Inc.) over approximately 1 × 10⁵ fibroblasts or adipocytes, or no feeder layer in 24-well plates. ALL cells and adipocytes were cultured with daunorubicin for various time intervals. ALL cell viability experiments were done in 96-well plates, using 0.75–1 × 10⁵ initial cells. In some experiments, BV173 cells were preloaded with daunorubicin for 1 hour at 37°C, pelleted, resuspended in ice-cold PBS, and plated on Transwells over either no feeder or adipocytes, and then collected at designated time points over the next 4 hours for flow cytometry.

All mouse experiments were approved by the Children's Hospital of Los Angeles (CHLA, Los Angeles, CA) Institutional Animal Care and Use Committee and performed in accordance with the U.S. Public Health Service Policy on Humane Care and Use of Laboratory Animals. C57BL/6J diet–induced obese and control mice (raised, respectively, on 60 kCal% or 10 kCal% fat diet from Research Diets) were purchased from The Jackson Laboratory. Obese and control male mice were used as a source of adipose tissue explants at 4–6 months of age. Mice were anesthetized with ketamine and xylazine, and intracardiac perfusion performed with PBS until liver clearing prior to harvesting of tissues. Adipose tissue was rinsed with cold PBS, cut into approximately 100-mg pieces, and washed twice with RPMI plus 10% FBS prior to culture in media with daunorubicin, For in vivo pharmacokinetic distribution, 3 obese and 3 control 13-week-old mice were injected with 5 mg/kg daunorubicin via tail vein. Mice were anesthetized 2 hours after daunorubicin injection as above, and blood samples collected via intracardiac puncture, followed by intracardiac perfusion as above. Blood, spleen, bone marrow, subcutaneous fat, and omental fat were collected for daunorubicin and daunorubicinol measurements using liquid chromatography/mass spectrometry (LC/MS; see below). Plasma was separated by centrifugation and white blood cells (WBC) were collected using Ficoll-Paque (GE Healthcare Life Sciences) according to manufacturer's protocol. Results from control and obese mice were combined as they were not qualitatively or statistically different.

All human samples were obtained and used after Institutional review board (IRB) approval and written informed consent and in accord with assurances filed with and approved by the U.S. Department of Health and Human Services. Subcutaneous abdominal adipose tissue biopsies were collected from a subset of obese adult female postmenopausal breast cancer survivors enrolled in an exercise intervention study 0–24 weeks out from completing chemotherapy and/or radiation (ref. 10; approved by the University of Southern California IRB; ClinicalTrials.gov: NCT01140282). Subjects were randomized to a supervised combined aerobic and resistance exercise program over 16 weeks or usual care, and underwent biopsy at baseline and after the intervention. Biopsies were collected as described previously (11). Adipose tissue biopsy samples were rinsed in normal saline, and then transported in saline on ice to CHLA (∼30 minutes), where they were immediately cut into approximately 100-mg sections. Some of these were cultured in complete media for 24 hours, and then fresh media was added with daunorubicin for experiments. While explant weight was closely matched between experiments, adipocyte number and viability were not assessed on fresh mouse or human tissues.

Bone marrow biopsy specimens were obtained from children aged 10–21 diagnosed with high-risk ALL, as described previously (12), under approval of the CHLA IRB (ClinicalTrials.gov: NCT01317940). Biopsies from day 29 (postinduction) were examined.

FACS analysis was done using a FACScan (BD Biosciences). Intracellular daunorubicin was measured using the phycoerythrin (PE) channel, taking advantage of the natural fluorescence of daunorubicin. MDR-1 surface expression was quantified using an allophycocyanin (APC)-conjugated anti-human MDR-1 antibody from BioLegend according to manufacturer's instructions. Cells stained with an APC isotype control antibody were used as a negative control. DAPI was used to distinguish live cells. For all samples, 1–5 × 10⁴ events were collected.

Adipocytes were grown and differentiated on poly-d-lysine–coated coverslips for analysis. These coverslips could then be placed in the bottom chambers of the Transwell cocultures. Daunorubicin fluorescence images were acquired with an LSM 700 confocal system mounted on an AxioObserver.Z1 microscope equipped with a 63×/1.4 Plan-APOCHROMAT objective lens and controlled with ZEN 2009 software (Carl Zeiss Microscopy). A 488-nm laser with a 560-nm long-pass filter was used for fluorescence excitation and emission. Transmitted laser light was collected to form a differential interference contrast (DIC) image simultaneously with the fluorescence image.

Paraformaldehyde-fixed bone marrow samples were embedded with paraffin. Samples were sectioned, mounted, and subjected to antigen retrieval with citrate buffer, pH 6.0, overnight. Endogenous peroxidases were inactivated with 0.3% H₂O₂. Nonspecific staining was blocked with 10% normal goat serum and 1% BSA before staining with polyclonal rabbit anti-human AKR1C1 (GeneTex), AKR1C2 (Cell Signaling Technology) or AKR1C3 (OriGene Technologies), and detected with polymerized peroxidase–labeled goat anti-rabbit immunoglobulin (Invitrogen; mouse adsorbed). The reaction was detected with 3,3′-diaminobenzidine (Millipore) and counterstained with Harris hematoxylin (Sigma). Images were acquired on a Leica DMI6000B Inverted Microscope (×40/1.25) with a Color CCD Digital Camera.

Adipose tissue was flash frozen and then lysed with QIAzol (Qiagen) using TissueLyzer II according to the manufacturer's protocol (Qiagen). qPCR was performed as described previously (Sheng and colleagues; ref. 9), with the following thermal profile: 10 minutes at 95°C followed by 40 repeats of 95°C for 15 seconds, 60°C for 1 minute, and a final dissociation stage of 95°C for 15 seconds, 60°C for 15 seconds, and 95°C for 15 seconds. See Supplementary Table S1 for primer sequences.

Human liver tissue lysate was from Abcam (ab29889). Total protein was extracted from adipocytes using lysis buffer [50 mmol/L Tris-HCl, 150 mmol/L NaCl, 0.1% SDS, 1% Nonidet I, 1 mmol/L phenylmethylsulfonylfluoride, 1% Halt Protease Inhibitor Cocktail (Thermo Scientific) and Phosphatase Inhibitor Cocktail Set II (Calbiochem)]. Adipose tissue was ground with lysis buffer using an electric rotator with glass pestle, followed by 20 strokes in a Dounce homogenizer. Lysates were sonicated with Bioruptor (Diagenode) for 10 minutes on ice and centrifuged for 15 minutes at 13,000 × g at 4°C. The supernatant was retained and protein concentration was quantified by BCA assay (Pierce Biotechnology). Proteins were separated using SDS-PAGE and transferred onto a nitrocellulose membrane using the iBlot 2 Dry Blotting System (Life Technologies). Membranes were blocked and then probed with specific primary antibodies, followed by horseradish peroxidase (HRP)-linked secondary antibodies. Bands were detected using SuperSignal West Pico Chemiluminescent Substrate (Thermo Scientific) and luminescence recorded with ImageQuant LAS 4000 (GE Healthcare Life Sciences).

Doxorubicin (50 μL at 1 μg/mL) was added as an internal standard (daunorubicin was the internal standard when doxorubicin and doxorubicinol were being analyzed). The entire sample was disrupted, and protein precipitated using 900 μL of ice-cold methanol and centrifuged at 13,000 rpm at 4°C for 5 minutes. The supernatant was isolated and evaporated to dryness. Cellular residues were reconstituted with 50-μL methanol with 0.1% formic acid, and 25 μL was injected into a Shimadzu Prominence HPLC linked to a Sciex API 3000. Each of the analytes was quantified using specific multiple reaction monitoring: 528.50→363.3, 530.60→321.30, 544.50→361.1, 546.5→363.2 and 445.63→98.60, for daunorubicin, daunorubicinol, doxorubicin, doxorubicinol, and mitoxantrone, respectively.

Activity was measured using a colorimetric assay based on ref. 13. Briefly, a reaction mixture containing 20 μmol/L of 9,10-Phenanthrenequinone (PQ; Sigma) and 200 μmol/L βNADPH (Sigma) was added to 1 μg of purified enzyme (rhAKR1C3 from R&D Systems, rhCBR1 from MyBioSource) or 5 μg lysates (mouse 3T3-L1 adipocytes, human breast adipose tissue) in sodium phosphate buffer (0.1 mol/L, pH 6.0) in a final volume of 100 μL/well. Stoichiometry of the reactions was determined by monitoring the decrease in NADPH/H⁺ absorbance at 340 nm for up to 30 minutes. Specific activity (pmol/min/μg) was determined using blank-adjusted OD/minute slope. In some experiments, indomethacin or luteolin (both 100 μmol/L final concentration) were added to the enzymes or lysates 5 minutes prior to adding the reaction mixture.

To determine whether human adipose tissue expresses enzymes known to metabolize anthracyclines, we evaluated four publicly available gene expression profile datasets (14–17), which included Affymetrix analyses of human subcutaneous and visceral adipose tissue from children and adults. Ranks for AKR and CBR genes were for analyzed each sample, independent of tissue source or clinical subgroup. When more than one detection probe was assigned to a gene, the one with the highest ranks were recorded. To investigate whether human adipose tissue expresses these enzymes at the protein level, we searched ProteomicsDB (18), a publicly available, mass spectrometry–based database of the human proteome. Log₁₀-normalized protein expression of AKR/CBR enzymes in tissues was used to generate a heatmap using Microsoft Excel.

Data are shown as mean ± SD. All experiments were performed at least three times. For flow cytometry of intracellular daunorubicin and surface MDR-1 expression, median fluorescence intensity (MFI) was reported. Two-sided, paired Student t tests were used to compare differences between the experimental conditions. A P value of less than 0.05 was considered statistically significant.

We previously showed that adipocytes protect murine leukemia cells from daunorubicin in both direct contact (7), and when separated by semipermeable membranes (8). To determine whether this ALL drug resistance is a consequence of decreased daunorubicin concentration in leukemic cells, we cultured human ALL cells for 24–48 hours in Transwells over no feeder or ∼1 × 10⁵ adipocytes. The presence of adipocytes significantly reduced daunorubicin accumulation in ALL cells, measured by fluorescence (Fig. 1A). To control for total number of cells in each well, additional Transwells with ALL cells over approximately 1 × 10⁵ 3T3-L1 fibroblasts were used, and showed that fibroblasts did not affect ALL cell daunorubicin concentration. The addition of adipocytes did not alter the surface expression of MDR-1 found on ALL cells, suggesting that reduced intracellular daunorubicin in ALL was not the consequence of increased cellular efflux (Fig. 1B). To confirm this, BV173 cells were preloaded with 18 μmol/L daunorubicin for 1 hour and plated alone or over adipocytes; the presence of adipocytes did not alter daunorubicin efflux from ALL cells (Fig. 1C and D).

To test whether adipocytes sequester daunorubicin from the media, 3T3-L1 adipocytes were cultured with daunorubicin for 4 hours. Adipocytes accumulated daunorubicin (measured by fluorescence) in a concentration-dependent manner, with signal visible primarily in the cytoplasm (Fig. 2A). To investigate whether this adipocyte sequestration would reduce media daunorubicin cytotoxicity, we preincubated media with 100 nmol/L daunorubicin (the EC₉₀ dose for BV173) with no feeder cells, fibroblasts, or adipocytes. ALL cells survived and proliferated better in media that had been preincubated with adipocytes when compared with fibroblasts or no feeder cells (Fig. 2B). This ability of adipocytes to detoxify the media was detectible even with daunorubicin concentrations as high as 1,000 nmol/L, albeit with diminished efficacy (Fig. 2C). Lysates from adipocytes cultured in high daunorubicin concentrations were toxic to BV173 cells (Fig. 2D), confirming that adipocytes did indeed sequester daunorubicin from the media in sufficient quantities to allow ALL cells to resist the daunorubicin cytotoxicity. Daunorubicin did not induce any significant morphologic changes or cell death in adipocytes by microscopy (not shown).

To determine whether adipocytes sequestered other anthracyclines and related drugs, we incubated 3T3-L1 adipocytes with various concentrations of doxorubicin and mitoxantrone for 16 hours. Adipocytes decreased the concentrations of both doxorubicin (Fig. 2E) and mitoxantrone (Fig. 2F) in the media, measured by LC/MS.

While adipocytes detoxified very high concentrations of daunorubicin from the media, their lysates were not as toxic as one would expect based on accumulation alone. We therefore hypothesized that adipocytes were metabolizing daunorubicin. As fluorescence cannot differentiate daunorubicin from its major metabolite, daunorubicinol, we used LC/MS to quantify both compounds. When cultured in daunorubicin for 16 hours, BV173 ALL cells accumulated daunorubicin, with nearly undetectable concentrations of daunorubicinol (Fig. 3A). However, when adipocytes were present, ALL cell daunorubicin concentrations were only approximately one-third as high.

Adipocytes took up daunorubicin in culture, but interestingly accumulated higher concentrations of daunorubicinol, a less cytotoxic metabolite (Fig. 3B). This suggests that adipocytes can deactivate daunorubicin. This finding was affirmed using human MSC-derived adipocytes, which also showed intracellular accumulation of daunorubicin and daunorubicinol (not shown). Furthermore, adipocytes rapidly reduced media daunorubicin concentration (Fig. 3C), while increasing media daunorubicinol concentration (Fig. 3D). Both of these effects were greater than observed in undifferentiated 3T3-L1 fibroblasts.

To test whether intact adipose tissue would sequester and metabolize daunorubicin, we incubated mouse adipose tissue explants in daunorubicin. After 16 hours, adipose tissue removed daunorubicin from the media, replacing much of it with released daunorubicinol (Fig. 4A). Adipose tissue accumulated both the cytotoxic daunorubicin and the inactivated daunorubicinol, demonstrating that intact adipose tissue can sequester, metabolize, and inactivate daunorubicin (Fig. 4B). This experiment was performed with adipose tissue from various anatomic sites, and demonstrated that adipose from all depots can efficiently metabolize daunorubicin to daunorubicinol. Human subcutaneous adipose tissue biopsy specimens also accumulated and metabolized daunorubicin, albeit with a high degree of variability in these experiments (Fig. 4C).

Although a comprehensive pharmacokinetic experiment was beyond the scope of this study, we tested whether adipose tissue would accumulate and metabolize daunorubicin in vivo. Two hours after a daunorubicin injection, plasma daunorubicin and daunorubicinol reached similar concentrations (Fig. 4D). While daunorubicin was detectible in the spleen, bone marrow, and circulating WBCs, little to no daunorubicinol was detected in these cells. In contrast, adipose tissue accumulated both daunorubicin and daunorubicinol. The ratio of daunorubicinol to daunorubicin was 0.60 ± 0.26 and 0.55 ± 0.21 in subcutaneous and omental adipose tissue, respectively. In contrast, this ratio was significantly lower in WBC (0.16 ± 0.11, P < 0.001 vs. both adipose tissues), and undetectable in spleen and marrow. These findings suggest that adipose tissue actively converts daunorubicin to daunorubicinol in vivo.

There are a number of enzymes capable of converting daunorubicin to daunorubicinol (19, 20), and so we next evaluated whether human adipose tissue expresses these enzymes. Four publicly available gene expression profiles (14–17) showed that human adipose tissue expressed high levels of many of these enzymes, including aldo-keto reductase (AKR)1A1, AKR1B1, AKR1C1, AKR1C2, AKR1C3, AKR7A2, carbonyl reductase (CBR)1, and CBR3 (Fig. 5A). AKR1B10 and AKR1C4 were not expressed (not shown). qPCR of human adipose tissue biopsy specimens confirmed high gene expression of all of these metabolic enzymes relative to β-actin (Fig. 5B), as well as undetectable levels of AKR1B10 and 1C4. ProteomicsDB, a publicly available database of human cellular proteomics (18), showed that adipocytes express high protein levels of these enzymes (Fig. 5C). Of note, this database showed adipocytes to have the highest protein levels of AKR1C1, AKR1C2, and AKR1C3 of all noncancerous tissues evaluated. Western blots confirmed protein expression of these AKR and CBR enzymes, with the exception of CBR3, in human subcutaneous adipose tissue biopsies and the human adipocyte cell line, ChubS7 (Fig. 5D). Furthermore, AKR1C1, 1C2, and 1C3 were also shown to be present in the cytoplasm of bone marrow adipocytes in children being treated for ALL (Fig. 5E).

To further verify that adipose tissue has AKR activity, we utilized a colorimetric assay based on NADPH (13). Lysates of both subcutaneous human adipose tissue biopsies and murine 3T3-L1 adipocytes showed AKR activity (Fig. 6A). This AKR activity was inhibited by the AKR1C inhibitor, indomethacin, and the CBR1 inhibitor, luteolin, although the latter did not reach statistical significance (Fig. 6B).

In this study, we present the novel finding that adipocytes sequester and metabolize the anthracycline, daunorubicin. This is the first report, to our knowledge, showing that adipocytes can metabolize and inactivate a therapeutic agent. This metabolism of daunorubicin, as well as sequestration of other chemotherapies such as doxorubicin and mitoxantrone, could reduce the concentration of active drugs in adipocyte-rich microenvironments, such as adipose tissue, omentum, and bone marrow. This is of particular importance as during leukemia treatment, bone marrow exhibits substantial fat accumulation (21). Furthermore, cancer treatment induces large increases in whole body adiposity (22), and obesity itself has been associated with higher adipose tissue expression of some of these enzymes (23, 24). Together, these changes may contribute to local reduction of cytotoxic activity of chemotherapy, leading to emergence of drug-resistant tumor cells and risk for treatment failure. We highlight a new role of the adipocyte in the emergence of chemotherapy resistance in the tumor microenvironment.

Anthracyclines are broadly used in treatment regimens for a wide variety of cancers, including leukemia, lymphoma, ovarian, pancreatic, breast cancers, bone, and soft tissue cancers. This new finding that fat can sequester and deactivate cytotoxic chemotherapy has wide implications and may partially explain why obese patients have poorer clinical response when compared with their leaner counterparts. In addition, the AKR and CBR isoenzymes are highly expressed in adipocytes and are known to metabolize a wide range of drugs; thus, it is possible that adipocytes could impact the efficacies of other drugs in relevant microenvironments.

We noted that adipocytes were less efficient in metabolizing doxorubicin when compared with daunorubicin. This was an unexpected finding as these anthracyclines differ by only one hydroxyl group, and they are cleared by similar isoenzymes (19). Decreased doxorubicin metabolism by adipocytes may reflect differing enzyme affinity between the two anthracyclines. For example, AKR1A1 metabolizes daunorubicin but not doxorubicin (25). Both daunorubicin and doxorubicin are substrates for AKR1B10, AKR1C1, AKR1C3, AKR7A2, and CBR1, but these enzymes preferentially metabolize daunorubicin when compared with doxorubicin (19, 26). On the other hand, others have reported that AKR expression contributes to doxorubicin resistance in breast cancer cells, even in the absence of detectible doxorubicinol accumulation in these cells or media (20). Nonetheless, adipocytes accumulated daunorubicin, doxorubicin, and mitoxantrone, suggesting that adipocytes can sequester all of these chemotherapies from their microenvironment, with differences only in subsequent intracellular metabolism.

While the rapid uptake and efficient deactivation of daunorubicin by adipocytes reduces daunorubicin concentration in the microenvironment and in nearby leukemia cells, the mass effect is insufficient to clearly alter the drug's plasma pharmacokinetics. Because the vast majority of anthracycline clearance from plasma occurs in the liver and kidney, peripheral adipocyte sequestration and metabolism would not be expected to significantly alter the plasma profile. This partially explains why Thompson and colleagues found that neither body mass index nor body fat were significantly correlated with doxorubicin or daunorubicin plasma clearance in children (27, 28). In fact, adiposity might be associated with a decreased plasma anthracycline clearance or increased AUC in adults (for example, see ref. 29). However, treatment failures and relapses in leukemia are most often present in the bone marrow, where our results show that increased adiposity could reduce available anthracycline levels. As we have previously shown that ALL cells migrate into adipose tissue under the influence of the chemokine CXCL-12 (30), adipose tissue itself could be an unrecognized sanctuary site for leukemia cells, where anthracyclines are unable to reach therapeutic levels. Thus, adipocyte anthracycline sequestration and metabolism may contribute to survival of local leukemia clones within the bone marrow and adipocyte, thus increasing the risk of residual disease at the end of induction therapy and eventual relapse.

In addition to reducing active daunorubicin concentrations in the leukemia microenvironment, adipocyte-mediated daunorubicin metabolism could contribute to its major long-term toxicity, cardiotoxicity. While much less cytotoxic to ALL cells than the parent compound, daunorubicinol has a longer half-life in both plasma and cardiac tissue, and has been shown to disproportionately contribute to cardiac toxicity (31). Adipocyte sequestration and slow release of daunorubicinol could result in an increased plasma half-life of this metabolite. This line of thought is supported by a pharmacokinetic study showing that doxorubicinol plasma clearance was decreased in children with >30% body fat (27). Thus, this mechanism may contribute to the observed link between obesity and anthracycline-related cardiotoxicity observed in animal models (32) and childhood cancer survivors (33).

There are some limitations to be considered in the data presented. While we have shown that adipocytes metabolize daunorubicin in vivo, we have not tested whether this leads to lower active daunorubicin concentration in nearby cancer cells in vivo, nor whether obesity alters ALL cell daunorubicin concentrations in vivo. In addition, although we showed that adipocytes take up daunorubicin at a much higher rate than ALL cells and fibroblasts, we have not characterized these uptake kinetics. These are important future experiments that should be done to further characterize these effects and help confirm the clinical relevance of the current studies. Furthermore, we identified the presence of several anthracycline-metabolizing enzymes in adipocytes, but have not identified the individual contribution that each enzyme has on daunorubicin metabolism. This level of detail is important, given the large number of these enzymes and isoforms, and their roles in metabolizing hormones, medications, and toxins. Although we are not the first to show that adipocytes express some AKR and CBR enzymes (18), our findings that adipocytes express these enzymes at very high levels compared with other tissues (Fig. 4) and that fat cells sequester and detoxify anthracyclines are highly novel. Adipose may be an underappreciated metabolic tissue that can influence cancer outcomes by creating a sanctuary microenvironment for ALL cells. Our use of cell culture, mouse experiments, and human tissues strengthen the veracity and clinical relevance of these findings.

In conclusion, this is the first report demonstrating that adipocytes sequester and efficiently metabolize a pharmaceutical agent. Specifically, adipocytes metabolize daunorubicin to a less toxic metabolite, and allow nearby ALL cells to evade daunorubicin-induced cytotoxicity. This finding could help explain why obese cancer patients are at risk of having a poorer outcome. Pharmacokinetic studies specifically in the tumor microenvironment will be necessary to determine the precise impact of adiposity on anthracycline-based treatment and efficacy in patients with ALL and other cancers.

No potential conflicts of interest were disclosed.

Conception and design: X. Sheng, S.G. Louie, S.D. Mittelman

Development of methodology: X. Sheng, J.-H. Parmentier, J. Tucci, O. Cortez-Toledo, S.G. Louie, S.D. Mittelman

Acquisition of data (provided animals, acquired and managed patients, provided facilities, etc.): X. Sheng, J.-H. Parmentier, J. Tucci, C.M. Dieli-Conwright, E. Orgel, S.G. Louie

Analysis and interpretation of data (e.g., statistical analysis, biostatistics, computational analysis): X. Sheng, J.-H. Parmentier, O. Cortez-Toledo, M.J. Oberley, M. Neely, E. Orgel, S.G. Louie, S.D. Mittelman

Writing, review, and/or revision of the manuscript: X. Sheng, J. Tucci, O. Cortez-Toledo, C.M. Dieli-Conwright, M. Neely, E. Orgel, S.G. Louie, S.D. Mittelman

Administrative, technical, or material support (i.e., reporting or organizing data, constructing databases): X. Sheng, J.-H. Parmentier, H. Pei

Study supervision: S.G. Louie, S.D. Mittelman

The authors thank Dr. Esteban Fernandez for assistance in the CHLA Imaging Core, and Dr. Michael Sheard and Tsen-Yin (Jackie) Lin in the CHLA FACS Core.

This work was supported by the National Cancer Institute at the NIH (R01 CA201444 and CA213129; to S.D. Mittelman), The Saban Research Institute (to S.D. Mittelman), The Office of the Dean of the Keck School of Medicine (to S.D. Mittelman), a Translational Research Program Award from the Leukemia & Lymphoma Society (to S.D. Mittelman and E. Orgel), The V Foundation for Cancer Research (to S.D. Mittelman), and the National Center for Advancing Translational Science of the NIH (UL1 TR001855 and UL1 TR000130; to C.M. Dieli-Conwright).

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