Blocking PGE2-induced tumour repopulation abrogates bladder cancer chemoresistance

Abstract

Cytotoxic chemotherapy is effective in debulking tumour masses initially; however, in some patients tumours become progressively unresponsive after multiple treatment cycles. Previous studies have demonstrated that cancer stem cells (CSCs) are selectively enriched after chemotherapy through enhanced survival. Here we reveal a new mechanism by which bladder CSCs actively contribute to therapeutic resistance via an unexpected proliferative response to repopulate residual tumours between chemotherapy cycles, using human bladder cancer xenografts. Further analyses demonstrate the recruitment of a quiescent label-retaining pool of CSCs into cell division in response to chemotherapy-induced damages, similar to mobilization of normal stem cells during wound repair. While chemotherapy effectively induces apoptosis, associated prostaglandin E2 (PGE2) release paradoxically promotes neighbouring CSC repopulation. This repopulation can be abrogated by a PGE2-neutralizing antibody and celecoxib drug-mediated blockade of PGE2 signalling. In vivo administration of the cyclooxygenase-2 (COX2) inhibitor celecoxib effectively abolishes a PGE2- and COX2-mediated wound response gene signature, and attenuates progressive manifestation of chemoresistance in xenograft tumours, including primary xenografts derived from a patient who was resistant to chemotherapy. Collectively, these findings uncover a new underlying mechanism that models the progressive development of clinical chemoresistance, and implicate an adjunctive therapy to enhance chemotherapeutic response of bladder urothelial carcinomas by abrogating early tumour repopulation.

Extended Data Figure 1. Human bladder urothelial carcinomas and their CK14 status pre- and post-neoadjuvant chemotherapy

a, Table summarizing the clinical information of 15 patients with bladder urothelial carcinomas, with paired pre- and post-chemotherapy tissues (n = 15). PDX-3 was derived from cancer tissues from patient 3 (bold). b, Patient subgroups showing an enrichment (increase) or persistence (maintain high expression) (left) or a resolution (absence) (right) of CK14 staining after chemotherapy treatment. c, d, Representative images of CK14 immunohistochemistry staining (original magnification, ×20), sub-classified as enrichment/ persistence (c) and resolution (d) of CK14 staining in matching bladder urothelial carcinoma tissues obtained pre- and post-neoadjuvant chemotherapy. Representative staining patterns of CK14 high-infiltrative staining (>25%), CK14 low-infiltrative staining (<25%), CK14 basal-restricted staining, and CK14 resolution are individually indicated.

Extended Data Figure 2. Cytotoxic chemotherapy induces CK14⁺ CSC proliferation despite reducing tumour size

a, Relative change in xenograft tumour volume from multiple xenograft tumour lines in response to GC chemotherapy or vehicle treatment (n = 6 per group). Xenograft tumours were derived from primary urothelial carcinoma patients (PDX-1 and PDX-2). b, Quantification of the percentage of CK14⁺ cancer cells in chemotherapy-treated and vehicle-treated xenograft tumours. c, Representative images demonstrating immunofluorescence staining of CK14⁺ cancer cells in chemotherapy-treated and vehicle-treated xenograft tumours. d, Representative images demonstrating immunofluorescence staining of pHH3 (red) and CK14 (green) in chemotherapy-treated and vehicle-treated xenograft tumours. Yellow arrows indicate CK14⁻ pHH3⁺ cells, white arrows indicate CK14⁺pHH3⁺ cancer cells. e, Graph quantifying the change in phospho-histone H3 positive (pHH3⁺) proliferating cells within CK14⁺ cancer cells (n = 6 per group). All data represent mean ± s.e.m. Box plots in b show twenty-fifth to seventy-fifth percentiles, with line indicating the median and whiskers indicating the smallest and the largest values. *P <0.05; **P <0.01; ***P <0.001 (two-tailed Student’s t-test). Scale bar, 100 μm.

Extended Data Figure 3. Methodology to purify CK14⁺cells for functional evaluation of sphere-forming and tumorigenic properties

a, Generation of a reporter construct to isolate viable CK14⁺ bladder cancer cells by FACS. A previously reported and validated human KRT14 gene promoter fragment was subcloned into a promoterless lentiviral construct that encodes a red fluorescent protein, DD-tdTomato. b–d, T24 high-grade urothelial carcinoma cells were stably transduced with the KRT14 reporter construct. FACS analysis validated that Tm⁺ CK14⁺ cells (red) represent a subpopulation of previously reported CD44⁺CD49f⁺ tumorigenic cells. e, Purified Tm⁺ CK14⁺ cancer cells demonstrated greater sphere-forming ability than Tm⁻ CK14⁻cancer cells in vitro (biological duplicates). f, Tm⁺ CK14⁺ bladder cancer cells are approximately 60-fold enriched for tumorigenic cells when engrafted in immunocompromised mice in vivo. Summary of tumour engraftment efficiency and image demonstrating tumour size after transplantation of 10, 50, 500 and 5,000 Tm⁺ CK14⁺ or Tm⁻ CK14⁻ cancer cells as purified by FACS. Data represent mean and range (e) and mean ± s.e.m (f). ***P <0.001 (two-tailed Student’s t-test).

Extended Data Figure 4. Cell viability of purified Tm⁺CK14⁺ and Tm⁻CK14⁻ cancer cells after GC chemotherapy treatment in vitro (raw FACS data for Fig. 1h)

Dot plots depict FACS analyses showing cell viability of Tm⁺ CK14⁺ (red) and Tm⁻ CK14⁻ (black) cancer cells following 11 consecutive days of chemotherapy treatment in vitro. The percentage of viable cells defined as annexin V⁻ PI⁻ is shown in the bottom left quadrant of each plot. Experiments were performed in biological duplicates.

Extended Data Figure 5. Cell cycle profiles of purified Tm⁺CK14⁺ and Tm⁻CK14⁻ cancer cells after GC chemotherapy in vitro (raw FACS data for Fig. 1i)

Histogram plots depict original FACS analyses of cell cycle profiles from Tm⁺ CK14⁺ (red) and Tm⁻ CK14⁻ (black) cancer cells after 11 consecutive days of chemotherapy treatment in vitro. Experiment was performed in biological duplicates. PI, propidium iodide.

Extended Data Figure 6. LRCCs are mutually exclusive to active proliferative cancer cells

a, b, Immunofluorescence staining to locate LRCCs (green, IdU⁺) at 0, 4 and 8 weeks of chase periods in patient-derived urothelial carcinoma xenograft (PDX-1) (a) and xenograft established from T24 high-grade urothelial carcinoma cells (b). c, d, Bar graph quantifying the percentage of LRCCs in patient-derived xenograft (PDX-1) (c) and immortalized cancer xenograft (T24) (d) at various chase periods (n = 4). e–g, Immunofluorescence staining to evaluate the localization of LRCCs (green, IdU⁺) and proliferating cells (red) using CldU (e), proliferating cell nuclear antigen (PCNA; f) or phospho-histone H3 (pHH3; g) in high-grade urothelial carcinoma (T24) at steady state. h, Immunofluorescence co-staining to locate IdU⁺ LRCCs, CldU⁺ proliferating and CK14⁺ cancer cells at various time points after GC chemotherapy. Data shown in c and d represent mean ± s.e.m. Scale bars, 100 μm.

Extended Data Figure 7. Celecoxib abrogates CK14⁺ cancer cell enrichment after GC chemotherapy in T24 and PDX-3 xenografts

a, In vivo treatment protocol recapitulating clinical regimen of one chemotherapy cycle in the presence or absence of celecoxib treatment. b, Immunofluorescence staining examining the percentage of CK14⁺ cancer cells in representative T24 xenograft tumours from various treatment groups. Scale bars, 1,000 μm. c, Immunohistochemical staining examining the percentage of CK14⁺ cancer cells in representative PDX-3 xenograft tumours from various treatment groups. Scale bars, 1,000 μm. Images in b and c are representative of n = 6 tumours analysed for each treatment group.

Extended Data Figure 8. Co-administration of aspirin diminishes thrombosis without impairing adjuvant effect of celecoxib

a, Antithrombotic effect of aspirin measurable by tail bleeding time. b, Temporal percentage change in tumour size after two cycles of celecoxib plus GC combination chemotherapy in the presence or absence of aspirin (n = 12 per group). c, Dynamics of COX1 and COX2 expression after GC chemotherapy, data shown for T24 cancer cells (displayed are representative blots from n = 3 experiments). All data represent mean ± s.e.m., tumour volume measurements shown in b are relative to mean tumour volume at day 0. *P <0.05; **P <0.01; ***P <0.001 (one-way ANOVA followed by Dunnett’s test for multiple comparisons); ns, not significant.

Extended Data Figure 9. Enrichment of the ‘wound-response gene signature’ in chemoresistant bladder urothelial carcinomas

a, GSEA validated an enrichment of the ‘wound-response gene signature’ (GO:0009611) in a panel of non-responding (or chemoresistant) human urothelial bladder carcinomas (n = 20; GSE48277), by comparing post-chemotherapy to pre-chemotherapy cancer tissues. Heat map demonstrates part of the genes within leading edge, including the COX2 gene PTGS2. b, Enlarged heat map for Fig. 4g. c, Enlarged heat map for Fig. 4h. d, Bioinformatics analysis of chemoresistant cancers in a panel of non-responding (or chemoresistant) human urothelial bladder carcinomas (n = 20; GSE48277) validated a significant increase of PTGS2 in post-chemotherapy tissues in comparison to matching pre-chemotherapy tissues. e, Dot plots representing the scoring of COX2 staining in bladder urothelial carcinoma tissues obtained pre-neoadjuvant chemotherapy in two subgroups of patients with different response to neoadjuvant chemotherapy (subgroups described in Extended Data Fig. 1a–c).

Extended Data Figure 10. Schematic model: recurrent CSC repopulation and its manifestation of chemoresistance

Cytotoxic chemotherapy effectively induces apoptosis but paradoxically elicits a wound response of bladder cancer stem cells to proliferate and repopulate residual tumours. Release of PGE₂ from neighbouring apoptotic cancer cells is sufficient to promote this CSC repopulation. In vivo administration of celecoxib effectively abolishes this PGE₂/COX2-mediated wound response gene signature, and attenuates progressive manifestation of chemoresistance in preclinical models of human urothelial carcinomas.

Figure 1. Cytotoxic chemotherapy induces CK14⁺cancer cell proliferation despite reducing tumour size

a, Kaplan–Meier analysis of bladder-cancer patients with various CK14 staining patterns (enrich/pers denotes an enrichment (increase) or persistence of CK14 staining; resolution denotes an absence) after neoadjuvant chemotherapy (n = 15). b, Relative change in xenograft tumour size after chemotherapy (n = 6 xenograft tumours per group). PDX-3, patient-derived xenograft from patient 3; T24, human bladder cancer cell line-derived xenograft. c, d, Representative immunofluorescence staining (c) and box plots quantifying the percentage of CK14⁺ bladder cancer cells after chemotherapy (d). DAPI, 4′,6-diamidino-2-phenylindole. e, Schematic demonstrating the transduction of hK14.tdTomato lentiviral reporter into urothelial carcinoma cells. f, Immunofluorescence staining verifying the specific expression of tdTomato (Tm)-positive signal in CK14⁺ cancer cells. g, Quantitative PCR (qPCR) analysis of KRT14 and UPK1B in cancer cell subpopulations in biological duplicates. h, i, Graphs summarizing viability (h) and corresponding percentage of S phase cells (i) in two cancer subpopulations after chemotherapy in biological duplicates. j, Cell cycle profiles for Tm⁺ CK14⁺ (red line) and Tm⁻ CK14⁻ (black line) cancer cells at indicated days after chemotherapy. PI, propidium iodide. Data in b represent mean ± s.e.m.; box plots in c show twenty-fifth to seventy-fifth percentiles, with line indicating the median, and whiskers indicating the smallest and largest values; data in g–i show mean and range. ***P <0.001 (log-rank test (a) and two-tailed Student’s t-test (b)). Scale bars, 100 μm.

Figure 2. Chemotherapy recruits CK14⁺ LRCCs to proliferate and divide

a, Experimental approach to label and localize LRCCs in bladder xenografts. Wk, week. b, Immunofluorescence staining to locate CK14⁺ cancer cells (red) that co-localize with LRCCs (green IdU⁺ cells) or proliferating cells (white CldU⁺ cells) at 8 weeks after IdU chase. c, Quantification of LRCCs within cancer subpopulations (n = 4 xenografted tumours). d, Immunofluorescence staining to evaluate the percentage and localization of LRCCs (green, IdU⁺), proliferating cells (red, CldU⁺) and dividing LRCCs (yellow, IdU⁺ CldU⁺) at various time after chemotherapy. e, Time kinetics revealing the percentage of LRCCs (green, IdU⁺), proliferating cells (red, CldU⁺), dividing LRCCs (yellow, IdU⁺ CldU⁺), and CK14⁺ cancer cells (white) at various time after chemotherapy (n = 4 xenografted tumours). All data represent mean ± s.e.m. *P <0.05, **P <0.01 (one-way analysis of variance (ANOVA) followed by Dunnett’s test for multiple comparisons). Scale bars, 50 μm.

Figure 3. PGE₂ release from neighbouring apoptotic cells induces CSC repopulation

a, Experimental approach to evaluate the role of PGE₂ release in modulating sphere-forming CSCs. b, FACS quantification of the percentage of viable cells at various time after chemotherapy (n = 3 experiments). c, Western blot analysing the correlative protein level of cleaved caspase-3 (CASP3) and COX2 at various time after chemotherapy (representative blot from n = 3). GAPDH was a loading control. d, Temporal release of PGE₂ by chemotherapy-treated cancer cells, measured by ELISA (n = 3 experiments). e, Immunofluorescence staining demonstrating relative localization of CASP3-positive apoptotic cells (white), PGE₂-positive signals (red) and CK14-positive cancer cells (green) in chemotherapy- or vehicle-treated xenograft tumours in vivo (n = 4 xenografted tumours). f, g, Relative fold changes in CASP3-positive (black), PGE₂-positive (red) and CK14-positive (green) cancer cells with time. h, Exogenous effects of dmPGE₂ on sphere-forming CSCs (n = 3 experiments). i, Inhibitory effects of a PGE₂-neutralizing antibody (Ab) and celecoxib in modulating sphere-forming CSCs (n = 4 biological replicates). All data represent mean ± s.e.m. *P <0.05, ***P <0.001 (Student’s t-test (h) and one-way ANOVA followed by Dunnett’s test for multiple comparisons (i)). Scale bars, 100 μm.

Figure 4. Celecoxib drug-mediated inhibition of PGE₂ pathway abrogates progressive development of chemoresistance

a, In vivo preclinical chemotherapy recapitulates clinical regimen with multiple treatment cycles and gap periods. b, Percentage change in tumour size after chemotherapy, in the presence or absence of celecoxib (n = 12 xenografted tumours (T24) per group). c, Representative haematoxylin and eosin images showing lung metastatic foci (asterisks) from various treatments. Scale bar, 500 μm. d, Quantification of lung metastatic foci across various treatments (n = 5 mice per group). e, Percentage change in tumour size of patient xenografts (PDX-3) after chemotherapy, in the presence or absence of celecoxib (n = 6 xenografted tumours per group). The four dashed boxes indicate the time frame of each GC chemotherapy cycle. f, Gene set enrichment analysis (GSEA) validated enrichment of ‘wound-response gene signature’ after chemotherapy in patients with bladder cancer classified as non-responders. NES, normalized enrichment score. g, GSEA revealing enrichment of wound-response gene signature in chemotherapy-treated bladder cancer xenografts. Heat map demonstrating genes within leading edge, including PTGS2 (encoding COX2), a core pathway component for PGE₂. h, GSEA and heat map highlighting abrogation of wound-response gene signature after celecoxib combination treatment. All data represent mean ± s.e.m. Tumour volume measurements are relative to mean tumour volume at day 0. *P <0.05, **P <0.01 (one-way ANOVA followed by Dunnett’s test for multiple comparisons).