The human pancreatic ductal adenocarcinoma cell lines, Panc-1 and Mia PaCa2, were obtained from The Cell Bank of Type Culture Collection of The Chinese Academy of Sciences and were routinely tested for mycoplasma before the experiments. The Panc-1 cells were cultured in DMEM (Gibco; Thermo Fisher Scientific) supplemented with 10% (v/v) FBS (Gibco; Thermo Fisher Scientific). Mia PaCa2 cells were maintained in DMEM supplemented with 10% (v/v) FBS and 2.5% (v/v) horse serum (Gibco; Thermo Fisher Scientific). The murine 3T3-L1 cell line is a well-established cell line that can be stably differentiated into mature adipocytes, and due to its good reproducibility, it is widely used for studies focusing on obesity, diabetes as well as the tumor microenvironment (14,16). Additionally, to the best of our knowledge, there are no stable human preadipocyte or adipocyte cell lines that can be used in such experiments. Thus, murine 3T3-L1 cells were chosen for the present study to ensure the stability of the coculture system, which has been widely used for studies on the crosstalk between cancer cells and adipocytes (14,17). Murine 3T3-L1 preadipocytes were obtained from the American Type Culture Collection and were maintained in DMEM supplemented with 10% newborn calf serum (Gibco; Thermo Fisher Scientific). All cells were cultured in a humidified incubator at 37°C supplied with 5% CO₂ air. A total of 2 days after reaching confluence, 3T3-L1 cell differentiation was induced by changing the medium to DMEM supplemented with 10% FBS (v/v), 1 µg/ml insulin, 0.5 mM 3-isobutyl-1-methylxanthine, and 1 µM dexamethasone for 2 days. The cells were then incubated in DMEM plus 10% (v/v) FBS and 1 µg/ml insulin for a further 2 days. Next, the differentiated mature adipocytes were cultured in DMEM supplemented with 10% (v/v) FBS. To study the crosstalk process between PC cells and adipocytes, the coculture model was constructed as previously described (11). Briefly, 8 days after induction, the mature adipocytes were cocultured with PC cells in a Transwell indirect coculture system (0.4 µm pore size; Corning, Inc.). Etomoxir (HY-50202, MedChemExpress) was added as a carnitine palmitoyl transferase 1 (CPT1) inhibitor, and CAY10499 (cat. no. 10007875, Cayman Chemical Company) was added as a nonselective lipase inhibitor.

The human tissues used in the present study were collected from a 78-year-old female patient with pancreatic ductal adenocarcinoma who underwent radical surgery in Huadong Hospital (Shanghai, China) in March 2019. Consent of the patient and approval from the Institutional Research Ethics Committee of Huadong Hospital, Fudan University (Shanghai, China; approval no. 2018K098) were obtained. Immunohistochemistry was performed as described previously (11). The paraffin-embedded tissues were stained with rabbit anti-FABP4 polyclonal antibody (pAb; cat. no. 12802-1-AP; ProteinTech Group, Inc.) overnight at 4°C. For each slide, representative images of adipocytes surrounding the normal tissue and adipocytes in the vicinity of the PC cells were obtained. Adipocyte cell sizes were assessed using ImageJ (version 1.8.0; National Institutes of Health).

To detect LDs in the 3T3-L1 adipocytes, the cells were cultured alone or with cancer cells for 5 days and then incubated in DMEM containing the BODIPY-493/503 lipid probe (0.1 µg/ml, cat. no. D3922, Invitrogen) for 15 min at room temperature. To detect LDs in PC cells treated with 200 µM oleic acid (OA: Beyotime, China), the cells were washed with PBS and fixed with 3.7% paraformaldehyde for 15 min at room temperature. The cells were then incubated with 0.1 µg/ml BODIPY and 5 µg/ml DAPI (Beyotime Institute of Biotechnology) for 15 min at room temperature. The stained cells were visualized using a confocal laser scanning microscope (Olympus) (magnification, ×200 or ×1,000).

Adipocytes were harvested after being cultured with or without PC cells for 5 days and then lysed with lysis buffer (Beyotime Institute of Biotechnology) for 15 min. The TG content of adipocytes was quantified using a TG assay kit (Applygen Technologies, Inc.). The total protein concentration was measured using a BCA assay (Thermo Fisher Scientific, Inc.), and the results are expressed as milligrams of TG per milligram of protein. The average of the control group was set as one, and all results are presented as the relative TG content.

The cells were harvested and lysed using TRIzol^® reagent (Invitrogen; Thermo Fisher Scientific, Inc.) according to the manufacturer's protocol. PrimeScript RT Master Mix (Takara Bio, Inc., cat. no. RR036A) was used to synthesize cDNA according to the manufacturer's protocol. Gene expression was determined using qPCR with a SYBR-Green PCR MasterMix Reagent (Applied Biosystems; Thermo Fisher Scientific, Inc.). The thermocycling protocol consisted of an initial denaturation step at 95°C for 10 min, followed by 40 cycles of 95°C for 15 sec and 60°C for 1 min. All amplifications and detections were performed using an ABI 7500 Real-Time PCR system (Applied Biosystems; Thermo Fisher Scientific, Inc.). Relative gene expression was calculated using the 2^−ΔΔCq method with 18S rRNA as an endogenous control (18). The average of the control group was set to one, and all results are presented as the relative mRNA expression. All assays were performed in triplicate. The primer sequences used in the present study are listed in Table I.

Total lysates were extracted using 2% SDS lysis buffer containing phosphatase and protease inhibitors (Roche Diagnostics GmbH). A BCA assay (Thermo Fisher Scientific, Inc.) was used to determine the protein concentration. A total of 20 µg protein was loaded per lane onto 12% SDS-gels, resolved using SDS-PAGE, transferred to 0.22 µm PVDF membranes (MilliporeSigma), and blocked for 1 h at room temperature with 5% skim milk. Each membrane was immunoblotted with the indicated primary antibodies at 4°C overnight and then incubated with a secondary antibody at 37°C for 1 h. Immunoreactive bands were visualized using an enhanced chemiluminescence detection kit (Thermo Fisher Scientific, Inc.) using an ImageQuant LAS 4000 System (GE Healthcare). The following primary and secondary antibodies were used in the present study: AKT (1:1,000; ProteinTech Group, Inc.; cat. no. 10176-2-AP), p-AKT^Ser473 (1:1,000, Cell Signaling Technology, Inc.; cat. no. CST4060S), HIF-1α (1:1,000; ProteinTech Group, Inc.; cat. no. 20960-1-AP), ANGPTL4 (1:1,000; ProteinTech Group, Inc.; cat. no. 18374-1-AP), GLUT1 (1:1,000; ProteinTech Group, Inc.; cat. no. 66290-1-lg), HK2 (1:1,000; ProteinTech Group, Inc.; cat. no. 22029-1-AP), LDHA (1:1,000; ProteinTech Group, Inc.; cat. no. 19987-1-AP), α-tubulin (1:1,000; ProteinTech Group, Inc.; cat. no. 66031-1-lg), p-AMPKα^Thr172 (1:1,000; Cell Signaling Technology, Inc.; cat. no. CST2535), AMPKα (1:1,000; ProteinTech Group, Inc.; cat. no. 10929-2-AP), p-STAT3^Tyr705 (1:1,000; Cell Signaling Technology, Inc.; cat. no. CST9145), STAT3 (1:1,000; Cell Signaling Technology, Inc.; CST12640), Snail (1:1,000; CST; CST9782), horseradish peroxidase-conjugated anti-rabbit IgG antibody (1:5,000; ProteinTech Group, Inc.; cat. no. SA00001-2) and goat anti-mouse IgG secondary antibody HRP conjugated (1:5,000; Signalway Antibody LLC; cat. no. L3032).

For the Transwell migration assay, 5×10⁴ Panc-1 or Mia PaCa2 cells in 200 µl DMEM were seeded into the upper chamber of a Transwell chamber with an 8-µm pore membrane (24-well insert; Corning, Inc.). The lower chamber was filled with media supplemented with 10% FBS, while the upper chamber contained serum-free media. Cells were allowed to adhere for 2 h prior to drug treatment and then incubated for 24 h for the migration assays. The cells that had not migrated through the pores were removed using cotton swabs, and the cells on the bottom of the membrane were fixed with 100% methanol and stained with 0.1% crystal violet at room temperature for 30 min. For the invasion assays, the Transwell chambers were coated in advance with 1 mg/ml Matrigel (cat. no. 356231, Corning, Inc.). Cancer cells that invaded through the Matrigel to the underside of the filter were stained with crystal violet at room temperature for 30 min. The number of migrated or invaded cells was counted in three randomly selected fields of view under a brightfield microscope (IX71; Olympus Corporation) (magnification, ×100). ImageJ was used to count the number of cells that had migrated or invaded in each field of view.

After PC cells were either cultured alone or cocultured with adipocytes for 5 days, the conditional medium was collected and centrifuged at 1,000 × g for 5 min at room temperature to remove debris. The supernatants were then stored at −80°C until required. Lactate released into the medium was measured using the Amplite™ Colorimetric L-Lactate Assay Kit (cat. no. 13815, AAT Bioquest) according to the manufacturer's instructions. Briefly, 50 µl L-Lactate standards and test samples were placed in a white, clear, bottom 96-well microplate. 50 µl L-Lactate working solution was then added to each well of the L-Lactate standard, blank control, and test samples to a final volume of 100 µl/well. The reaction was incubated at room temperature for 2 h in the dark. The absorbance was measured at 575/605 nm. A standard curve based on the absorbance of the L-lactate standards was drawn, and the lactate concentrations in the supernatants were then calculated.

PC cells were cultured with or without adipocytes for 5 days. The cells were then collected by centrifugation at 1,000 × g for 10 min at room temperature and immediately fixed in 2.5% glutaraldehyde, 4% paraformaldehyde, and 0.002% picric acid in a 0.1 M (pH 7.3) cacodylate buffer at 4°C for 3 h. Tissue slices were then postfixed in 1% OsO₄ in the same buffer at 4°C for 3 h, dehydrated in a graded acetone series, and embedded in Epon resin. For electron microscopy, 70-nm thick sections were cut from tissue resin blocks. The sections were then transferred to formvar-coated copper mesh grids and double-stained with saturated uranyl acetate for 30 min, followed by lead citrate for 15 min at room temperature. The ultra-thin sections on the grids were examined in a JEOL JEM-1400 plus transmission electron microscope at 80 kV. ImageJ (version 1.8.0; National Institutes of Health) was used to count the number of LDs.

After culturing with or without adipocytes for 5 days, PC cells were seeded into 96-well plates at a density of 40,000 cells/well and incubated overnight. Mitochondrial function and cellular glycolytic capacity were determined using the Seahorse Bioscience XF96 Extracellular Flux Analyzer (Seahorse Bioscience) and a Seahorse XF Glycolysis Stress Test Kit and a Cell Mito Stress Test Kit, according to the manufacturer's protocol. For ECAR assessment, cells were incubated under basal conditions with non-buffered RPMI 1640 followed by sequential injection of 10 mM glucose and 1 mM mitochondrial poison (oligomycin; MilliporeSigma). OCR was evaluated under basal conditions, followed by a sequential injection of 1 µM oligomycin, 1 µM fluoro-carbonyl cyanide phenylhydrazone (FCCP; MilliporeSigma), and 2 mM antimycin A and rotenone (MilliporeSigma). Both ECAR and OCR measurements were standardized to total protein content.

RNA-seq was performed on 3T3-L1 cells cultured with or without Panc-1 PC cells for 5 days as previously described (11). The sequencing data have been deposited in the Gene Expression Omnibus under accession number GSE123939. GSEA was performed using GSEA software (http://www.broadinstitute.org/gsea/index.jsp). A gene set enrichment score (ES) estimating genes from a predefined gene set was calculated using GSEA. The thresholds for significance were set by permutation analysis with 1000 gene-set permutations.

Panc-1 PC cells were cultured with or without mature adipocytes for 5 days and then resuspended in an ~8-fold volume of lysis buffer (4% SDS, 100 mM HEPES, pH=7.6) containing a protease inhibitor cocktail and PMSF. The homogenate was sonicated on ice for 30 min. The sample was centrifuged at 25,000 × g for 30 min at 4°C, and the supernatant was stored at −80°C. Proteins were reduced in 10 mM DTT for 1 h at 37°C. Protein samples were then cooled to room temperature (RT). The cysteines were blocked in darkness with 30 mM IAA at 37°C for 30 min. The extracted protein was mixed in equal amounts according to the groups and then precipitated overnight in acetone. The protein samples were then resuspended in 1 M urea buffer and digested with trypsin overnight. Next, peptides were isotopically labeled with iTRAQ reagents (Applied Biosystems; Thermo Fisher Scientific, Inc.) for 2 h at RT. The labeling reaction was then stopped using water. The samples were then separated and identified using a TripleTOF 4600 mass spectrometer (Applied Biosystems; Thermo Fisher Scientific, Inc.).

All statistical analysis was performed using GraphPad Prism version 5.0 (GraphPad Software, Inc.). All results are presented as the mean ± standard deviation of at least three repeats. The differences between two groups were determined using an unpaired two-tailed Student's t-test. Multiple-group comparisons were conducted using one-way analysis of variance (ANOVA) followed by Tukey's post hoc test. P<0.05 was considered to indicate a statistically significant difference.

Although obesity adversely affects the long-term outcomes of patients with PC, the crosstalk between adipocytes and PC cells has not been fully elucidated (19). Immunohistochemical staining of the adipocyte marker FABP4 in human PC tissue demonstrated that adipocytes were present in the TME (Fig. 1A). It was also found that adipocytes directly adjacent to the tumor were smaller in size compared to those further away (P<0.0001; Fig. 1B). To assess the role of adipocytes in PC progression, an adipocyte-PC cell coculture system was established (Fig. 1C). Our previous study revealed the significant phenotypic alterations in mature adipocytes induced by pancreatic Panc-1 and MIA PaCa2 cells (11); in the present study, the relationship between adipocytes and PC cells was further explored. 3T3-L1 preadipocytes were differentiated into mature adipocytes and then cocultured with human Panc-1 or MIA PaCa2 cells. It was found that cocultured Panc-1 cells exhibited an elongated mesenchymal morphology compared to monocultured Panc-1 cells, which showed a characteristic epithelial morphology and formed compacted colonies (Fig. 1D). It was also found that PC cells cocultured for 5 days with mature adipocytes had a significantly increased migratory and invasive capacity (P<0.05; Fig. 1E and F). Taken together, these data indicated that adipocytes surrounding tumors may have promoted PC progression.

To decipher the potential mechanisms responsible for the increased aggressiveness of cocultured cancer cells, mass spectrometry was used to analyze the protein contents of Panc-1 monocultures and those cocultured with adipocytes for 5 days. GSEA revealed a striking overrepresentation of hallmark database-defined pathways involved in hypoxic signaling in Panc-1 cells cocultured with adipocytes compared to those that were monocultured (Fig. 2A). The majority of the top ten upregulated proteins (Table II) are well-established downstream factors of hypoxic signaling, such as ANGPTL4, ENOG, and LDHA (20–22). Whole proteome bioinformatics analysis also revealed that certain metabolic processes and the epithelial-mesenchymal transition were activated in cocultured tumor cells (Fig. 2A). Based on the protein-protein interaction network associated with the enriched functional pathways of carbohydrate metabolism, tissue morphology, and cancer, it was further confirmed that HIF-1α was activated in the cocultured Panc-1 cells (Fig. 2B). Congruently, there was a robust increase in HIF-1α and its related downstream proteins (SNAIL, ANGPTL4, and glycolytic-associated proteins) in the cocultured tumor cells (Fig. 2C), which was further confirmed by RT-qPCR analysis (Fig. 2E and F). It was also found that lactate production was significantly increased in the cocultured cells (P<0.01; Fig. 2D). Together, these data suggest that adipocytes enhance HIF-1α signaling and can reprogram the tumor metabolic pattern to induce a shift towards anaerobic glycolysis in PC cells under in vitro coculture conditions.

Given the enhanced ability of glucose utilization in cancer cells, whether coculturing of cells resulted in altered glucose metabolism in adipocytes was next assessed. To address this, RNA-sequencing on mature 3T3-L1 adipocytes cultured alone or with Panc-1 PC cells for 5 days was performed. GSEA of differentially expressed transcripts in cocultured adipocytes compared with those cultured alone revealed a marked enrichment of gene sets corresponding to the insulin signaling pathway and the JAK-STAT3 pathway (Fig. 3A). In addition, there was a robust decrease in genes associated with the insulin signaling pathway in cocultured adipocytes (Fig. 3B), indicating insulin resistance in the adipocytes. It was also found that coculturing the adipocytes with cancer cells increased STAT3 phosphorylation (Fig. 3C). Additionally, the increased phosphorylation of STAT3 in adipocytes treated with 20 ng/ml IL-6 resulted in the downregulation of genes related to the insulin signaling pathway, such as GLUT4 and IRS1 (P<0.05; Fig. 3D and E). In humans, SOCS3 expression has been shown to be associated with the JAK-STAT3 pathway and insulin resistance (23). In line with this, significantly increased SOCS3 expression in adipocytes treated with IL-6 or cocultured with Panc-1 or MIA PaCa2 PC cells was observed (P<0.001; Fig. 3F). Given the close relationship between the JAK-STAT3-SOCS3 axis and the development of obesity-associated disorders, such as insulin resistance (24), it was hypothesized that PC cells could induce an insulin-resistant phenotype in adipocytes through the JAK-STAT3-SOCS3 axis.

Next, both the ECAR and OCR were measured in PC cells using a Seahorse assay. After monoculture or coculture with 3T3-L1 adipocytes for 5 days, the PC cells were digested and seeded into 96-well plates for further tests. It was found that the ECAR did not differ significantly between the two conditions, indicating no alteration in the rate of glycolysis in PC cells after coculture with adipocytes (Fig. 4A and B). In contrast, the cocultured cancer cells underwent significantly increased OCR in both basal and maximal-uncoupled states compared with monocultured cancer cells (Fig. 4C and D), suggesting enhanced fatty acid β-oxidation (FAO) in the cocultured cancer cells. In cancer, the JAK-STAT3 pathway regulates lipid metabolism through FAO (25), and AMPK favors energy-producing processes by activating β-oxidation (26). Congruently, the presence of mature adipocytes increased STAT3 and AMPK phosphorylation and decreased AKT phosphorylation (Fig. 4E). As adipocytes store LDs, it was hypothesized that adipocytes provide lipids to cancer cells to enhance their FAO ability. To explore the changes in lipids in cocultured PC cells, electron microscopy was performed. The results showed there was a substantial increase in the number of LDs in the cocultured Panc-1 and MIA PaCa2 cancer cells compared with that in the monocultured cells (P<0.05; Fig. 4F and G). Substantial changes in the mitochondrial ultrastructure in the cocultured MIA PaCa2 cells were also observed (Fig. 4G), suggesting enhanced respiratory chain activity in cancer cells after coculture with adipocytes. Taken together, these results showed that adipocytes resulted in increased lipid content and metabolic reprogramming in PC cells.

The above data showed that during coculture, adipocytes stimulated increased invasiveness and enhanced FAO in PC cells. Thus, whether the utilization of stored lipids in cancer cells was associated with tumor malignancy was further examined. Etomoxir can inhibit CPT1, which serves as the primary rate-limiting factor in the transport of fatty acids to the mitochondria.

First, cancer cells were cocultured with or without adipocytes for 5 days with the addition of etomoxir and then the cancer cells' invasive ability was assessed. It was found that the inhibition of FAO by etomoxir during coculture did not hamper the invasive ability of the cocultured cancer cells, in agreement with the unchanged OCR (Fig. 5A and B). As catabolism of stored LDs promotes cancer invasion and migration (27), whether excess stored lipids contributed to the increased invasive ability of cocultured PC cells compared with monocultures was assessed. Treating the cancer cells with etomoxir to inhibit FAO or with CAY10499 to inhibit lipolysis during the invasion assay significantly reduced the invasion of the cocultured PC cells (P<0.01; Fig. 5C and D). These data indicate that the accumulation of LDs during tumor cell coculture with adipocytes and the utilization of excess lipids during the process of tumor metastasis together resulted in the increased invasive ability of the cocultured PC cells. In agreement with the increased levels of LDs in cocultured cancer cells (Fig. 4F and G), it was also shown that the fatty acid transporter-related gene (FATP1) and lipid storage-related genes (DGAT1, DGAT2, GPAT4, and PLIN2) were upregulated in the cocultured PC cells (Fig. 6A and B), suggesting that coculture with adipocytes promoted fatty acid uptake and storage in the cancer cells. To further confirm this, Panc-1 and MIA PaCa2 cancer cells were first pretreated with exogenous OA, which resulted in lipid accumulation (Fig. 6C and D). Preloading exogenous LDs increased cancer cell aggressiveness, and this effect was abrogated by etomoxir or CAY10499 treatment (P<0.01; Fig. 6E and F). Taken together, these data demonstrated that the presence of adipocytes increased the metastatic capacity of the PC cells, and this was largely due to LD accumulation in the cocultured tumor cells.

The above data indicated that adipocytes altered PC cell lipid metabolism. Next, whether coculturing with cancer cells also influenced the lipid metabolism of adipocytes was assessed. First, a decrease in the size of LDs was found in cocultured adipocytes, which was consistent with the reduced TG content (P<0.01; Fig. 7A and B). The GSEA of transcripts that were downregulated in the cocultured adipocytes showed enrichment in gene sets corresponding to oxidative phosphorylation, adipogenesis, and fatty acid metabolism. Further analysis using Gene Ontology and biological process compilation confirmed that both fatty acid catabolic and anabolic processes were suppressed in cocultured adipocytes (Fig. 7C). The GSEA of transcripts that were downregulated in the cocultured adipocytes also revealed marked enrichment of gene sets associated with adipogenesis (Fig. 7D). In agreement with this, RT-qPCR analysis of the adipocytes revealed the reduced expression of key upstream genes of adipogenesis when cocultured with PC cells (Fig. 7E). Taken together, these results indicate that PC cells downregulated lipid metabolism and adipogenesis in the cocultured adipocytes.