Authors: Anubhav Dhull, Anunay James Pulukuri, Aqib Iqbal Dar, Nina Julia Palmer, Joan Castaneda Gonzalez, Anu Rani, Rishi Sharma, Clifford E. Berkman, Anjali Sharma
Categories: Article, PAMAM dendrimer, prostate cancer, targeted drug delivery, camptothecin, click chemistry
Source: ACS applied nano materials
Authors: Anubhav Dhull, Anunay James Pulukuri, Aqib Iqbal Dar, Nina Julia Palmer, Joan Castaneda Gonzalez, Anu Rani, Rishi Sharma, Clifford E. Berkman, Anjali Sharma
Prostate cancer (PC) remains a major global health challenge, particularly in its advanced, treatment-resistant form. Although Camptothecin (Campto) is a potent topoisomerase I inhibitor with strong antiproliferative and pro-apoptotic activity, its clinical utility is limited by poor aqueous solubility, instability, and systemic toxicity. To address these challenges, we developed a prostate-specific membrane antigen (PSMA)-targeted dendrimer-Camptothecin conjugate (PD-Campto-CTT1298) that improves drug solubility, enables receptor-mediated uptake, and enhances therapeutic effects. This platform was synthesized by conjugating Campto and a high-affinity PSMA ligand (CTT1298) to a generation-4 hydroxyl-terminated PAMAM dendrimer (PD) via copper-catalyzed and strain-promoted azide–alkyne cycloaddition reactions. The resulting conjugate exhibited high aqueous solubility, formulation stability, and efficient drug loading. In vitro drug release studies demonstrated pH- and esterase-responsive Campto release mimicking the tumor microenvironment. PD-Campto-CTT1298 showed selective uptake in PSMA-positive PC cells (PC3-PIP, C4–2B), leading to enhanced cytotoxicity, apoptosis induction, mitochondrial dysfunction, and reactive oxygen species generation. Confocal imaging and flow cytometry confirmed receptor-specific internalization via clathrin-mediated endocytosis. Moreover, PD-Campto-CTT1298 suppressed VEGF-A secretion and disrupted angiogenesis in HUVEC tube formation assays, indicating antiangiogenic activity. Combination therapy with Olaparib demonstrated synergistic effects. Overall, PD-Campto-CTT1298 represents a promising strategy for PSMA-targeted prodrug delivery, offering a multifaceted therapeutic approach through enhanced solubility, intracellular release, and PSMA-positive tumor-specific enhanced cytotoxicity.
Prostate cancer (PC) is one of the most prevalent malignancies in men worldwide, ranking as the second most frequently diagnosed cancer and the fifth leading cause of cancer-related deaths, with an estimated 1.46 million new cases and 396,000 deaths globally in 2022.^1,2^ This burden is expected to increase significantly in the coming decades. By 2040, projections estimate approximately 2.4 million new cases and 712,000 deaths, driven primarily by the aging and growing global population.^3^ Standard treatment options for PC include surgery, radiotherapy, chemotherapy, and hormone therapy.^4^ In recent years, therapeutic strategies have increasingly focused on targeting the androgen signaling pathway through androgen deprivation therapy (ADT).^5^ However, approximately 20–30% of patients experience tumor recurrence and progression to advanced castration-resistant PC (CRPC), transitioning from hormone-sensitive to metastatic disease.^6^ Despite the approval of several new agents, the optimal sequencing of therapies in advanced PC remains undefined.^7^
Chemotherapeutic drugs such as docetaxel, cabazitaxel, and mitoxantrone are widely used in advanced PC and offer clinical benefits including tumor regression and extended survival.^8^ However, their use is associated with significant toxicities, including cytopenia, nausea, neutropenic sepsis, and pancytopenia, which often lead to dose modifications and/or supportive interventions.^9,10^ Camptothecin (Campto) and its derivatives are also being evaluated in various clinical trials for cancer treatment due to their potent inhibition of DNA topoisomerase I, which disrupts DNA replication and induces apoptosis.^11^ Recent studies further indicate that Campto interferes with androgen receptor-mediated transcription, contributing to tumor growth suppression in PC.^12^ However, its clinical use is restricted by poor water solubility, rapid hydrolysis of the lactone ring, systemic toxicity, and acquired resistance.^13^ These limitations are particularly critical in PC, where drug access to tumor tissue can be further hindered by heterogeneous vascularization and a dense stromal microenvironment. Therefore, strategies that improve the solubility, stability, and tumor selectivity of Campto are needed for full therapeutic potential in this setting. To address these limitations, several Campto analogs such as rubitecan, exatecan, irinotecan, and topotecan have been developed.^14^ Some of these derivatives are still undergoing clinical trials, while others have been approved by the United States Food and Drug Administration (FDA) and are now used in clinical use.^14–16^ Notably, polymeric Campto-based prodrugs have shown promise in improving therapeutic outcomes and overcoming resistance in various cancers, including PC and ovarian cancer.^17,18^ Cyclodextrin-based Campto formulations such as EP0057 have demonstrated improved efficacy with reduced toxicity and are currently in clinical trials.^19^ These advancements underscore the potential of targeted delivery systems and prodrug strategies to enhance the clinical utility of Campto.^20^
Among various nanocarriers, dendrimers have emerged as attractive platforms for drug delivery and imaging due to their well-defined architecture, monodispersity, and multivalency, which facilitate the conjugation of drugs, targeting ligands, and imaging agents.^21–24^ Polyamidoamine (PAMAM) dendrimers (PD), in particular, have been extensively studied for biomedical applications, including cancer diagnosis and therapy.^25,26^ Their ability to enhance drug solubility and enable targeted delivery makes them promising candidates for clinical use.^27–29^ In our previous work, we developed a PC-targeting delivery platform (PD-CTT1298) combining PD with CTT1298, a high-affinity ligand for prostate-specific membrane antigen (PSMA).^30^ PSMA is a type II trans-membrane glycoprotein that is overexpressed on PC cells and exhibits receptor-mediated internalization, making it an ideal target for selective drug delivery.^31,32^ The CTT1298 ligand binds irreversibly to PSMA, facilitating highly specific uptake in prostate tumors.^33,34^ In a xenograft mouse model, PD-CTT1298 selectively targeted PSMA-positive prostate tumor demonstrating its potential for precise and effective drug delivery.^30^
Building on this targeted approach, in this study, we explore the combination of PD-CTT1298 with Campto as a strategy for improving the therapeutic outcomes of Campto in PC. We report the synthesis, characterization, and in vitro evaluation of a dual-functional PSMA-targeted dendrimer conjugated with Campto (PD-Campto-CTT1298). Using copper-catalyzed alkyne-azide click (CuAAC) and strain-promoted alkyne-azide click (SPAAC) chemistry, we generated a soluble, stable, and receptor-specific dendrimer-prodrug system designed for efficient delivery to PC cells. The therapeutic potential of PD-Campto-CTT1298 was evaluated through cellular uptake, cytotoxicity, mechanistic studies, and antiangiogenic assays in PSMA-expressing PC models. This approach integrates rational drug design with PC cell targeting, thereby offering a comprehensive strategy to overcome key challenges in PC chemotherapy.
All starting materials and reagents were obtained from Sigma-Aldrich and Merck. Analytical-grade reagents and solvents used in the synthesis were used as received. Reactions requiring anhydrous conditions were conducted under a positive nitrogen atmosphere, with all glassware dried in an oven beforehand. Analytical thin-layer chromatography (TLC) was performed on silica gel 60 F254 aluminum-backed plates, and spots were visualized using UV light or staining reagents. Compounds were purified by flash column chromatography on silica gel 60 (230–400 mesh). Porcine liver esterase (PLE) (16 U/mg) was purchased from Sigma-Aldrich. Dialysis membranes (Spectra/Por) were sourced from Repligen. DBCO-C6-CTT1298 was synthesized using -a previously reported procedure.^30^
Nuclear Magnetic Resonance (NMR) spectra were acquired using a Bruker 500 MHz high-resolution NMR spectrometer with samples prepared in deuterated solvents such as chloroform (CDCl3), deuterated DMSO (DMSO-d6), or deuterated water (D2O). Chemical shifts (δ) for ^1^H NMR are given in parts per million (ppm) and are referenced to the solvent peak. Coupling constants (J) are reported in hertz (Hz), with the following abbreviations used to describe signal s = singlet, d = doublet, t = triplet, q = quartet, and m = multiplet. MALDI-TOF spectra were obtained using electrospray ionization (ESI) with direct infusion on a Bruker MALDI-TOF (using a trans-2-[3-(4-tert-butylphenyl)-2-methyl-2-propenylidene] malononitrile matrix). High-resolution mass spectrometry (HRMS) spectra were acquired using a Waters Q-Tof Premier mass spectrometer operating in positive-ion, V-mode using electrospray ionization.
Particle size and zeta potential were measured by Dynamic Light Scattering (DLS) using a Malvern Zetasizer Nano 90 (Westborough, MA) at 25 °C. For particle size analysis, PD-Campto-CTT1298 was dissolved in deionized water (18.2 MΩ) to a final concentration of 0.5 mg/mL. The solution was filtered through 0.2 μm syringe filters (Pall Corporation, HT Tuffryn membrane, 0.2 μm) directly into a UV-transparent disposable cuvette (12.5 × 12.5 × 45 mm). For zeta potential measurements, a sample was prepared at a concentration of 0.2 mg/mL in 10 mM NaCl following the same procedure. Zeta potential analysis was performed in a Malvern Zetasizer Nanoseries disposable folded capillary cell.
High-performance liquid chromatography (HPLC) was used to assess the purity of small molecules as well as dendrimers and dendrimer-drug conjugates and to evaluate drug release studies. Analyses were performed on a Waters Acquity Arc system (Milford, MA, USA) equipped with binary pumps, a 2998 PDA detector, and a 2475 fluorescence detector using Waters Empower software. Samples were analyzed on a Waters C18 Symmetry 300 column (5 μm, 4.6 × 250 mm) with a gradient flow method. The gradient started at 10 (Solvent A, 0.1% TFA and 5% ACN in water; Solvent B, 0.1% TFA in ACN), ramped to 75 (A:B) over 25 min, then returned to 10 (A:B) at 35 min, with a flow rate of 1 mL/min. Dendrimers and drug conjugates were monitored at 210 nm (dendrimer absorbance), 254 nm (aromatic compound absorbance), and 369 nm (Campto absorbance). The cyanine 5 (Cy5) conjugates were also monitored at 650 nm (Cy5 absorbance).
To a stirring of solution of hexynoic acid (2) (483 mg, 1.5 equiv, 4.31 mmol) in anhydrous N,N-dimethylformamide (DMF; 7 mL) was added 1-ethyl-3-(3-(dimethylamino)-propyl)carbodiimide hydrochloride (EDC·HCl) (1.1 g, 2.0 equiv, 5.75 mmol) and stirred for 15 min. It was followed by the addition of Campto (1) (1 g, 1.0 equiv, 2.87 mmol) and 4-(dimethylamino)pyridine (DMAP) (175 mg, 0.5 equiv, 1.43 mmol). The reaction mixture was allowed to stir at room temperature for 12 h. The progress of the reaction was monitored with the help of thin-layer chromatography (TLC) in UV light. The reaction mixture was diluted with dichloromethane (DCM) and the organic layer was washed with water (5 × 30 mL) and brine (2 × 30 mL), dried (Na2SO4), filtered, and evaporated in vacuo. The crude product was purified by silica flash column chromatography [methanol/dichloromethane, 95 (v/v)] to afford compound 3 as a white solid in 73% yield.
^1^H NMR (500 MHz, DMSO-d6) δ 8.74 (s, 1H), 8.26–8.13 (m, 2H), 7.91 (t, J = 8.4 Hz, 1H), 7.76 (t, J = 7.4 Hz, 1H), 7.10 (s, 1H), 5.54 (d, J = 3.0 Hz, 2H), 5.34 (d, J = 3.5 Hz, 2H), 2.89 (s, 1H), 2.68 (t, J = 7.3 Hz, 2H), 2.33–2.14 (m, 4H), 1.77 (m, 2H), 0.96 (t, J = 7.4 Hz, 3H) (Figure S1).
^13^C NMR (125 MHz, DMSO-d6) δ: 172.10, 167.77, 157.05, 152.79, 148.32, 146.50, 145.92, 132.10, 130.93, 130.32, 129.40, 129.03, 128.48, 128.23, 119.25, 95.21, 83.99, 76.31, 72.37, 66.74, 50.75, 32.48, 30.65, 23.85, 17.36, 8.05 (Figure S2).
HPLC 95.26%, retention 20.572 min (Figure S3).
Mass: m/z: calculated for C26H22N2O5 [M + H]^+^: 443.1607; found 443.1588 (Figure S4).
Azido hexanoic acid (5) (143 mg, 13.0 equiv, 0.91 mmol) was dissolved in anhydrous DMF (5 mL), followed by the addition of EDC·HCl (324 mg, 24.0 equiv, 1.68 mmol). The mixture was stirred for 15 min at room temperature to activate the carboxylic acid functionality. This activated solution was then added dropwise to a separately prepared solution of PD (4) (1.0 g, 1.0 equiv, 0.07 mmol) in anhydrous DMF (5 mL) under continuous stirring. Subsequently, DMAP (46 mg, 6.0 equiv, 0.38 mmol) was added and the reaction was stirred at room temperature for 24 h. The reaction completion was confirmed by the shift in the retention time of the chromatogram in HPLC. The crude product was purified via dialysis using a 1 kDa molecular weight cutoff membrane against deionized water for 24 h. The aqueous solution was lyophilized to afford PD-azide (6) as a white solid in 84% yield.
^1^H NMR (500 MHz, DMSO-d6) δ 8.00–7.72 (m, 124H, Dendrimer-internal amide-H), 4.65 (s, 57H, Dendrimer–OH), 3.93 (t, J = 5.8 Hz, 26H, Dendrimer-Ester CH2), 3.40–3.17 (m, 250H, Dendrimer-CH2), 3.14–2.91 (m, 245H, Dendrimer-CH2), 2.67–2.50 (m, 248H, Dendrimer-CH2), 2.42–2.30 (m, 120H, Dendrimer-CH2), 2.25–2.21 (m, 26H, Linker-CH2), 2.21–2.06 (m, 248H, Dendrimer-CH2), 1.53–1.38 (m, 48H, Linker-CH2), 1.29–1.20 (m, 24H, Linker-CH2) (Figure S5).
HPLC 99.89%, retention 12.501 min (Figure S6).
To a stirred solution of compound 6 (300 mg, 1.0 equiv, 0.02 mmol) in DMF (1.5 mL) was added a solution of compound 3 (84.5 mg, 10.5 equiv, 0.19 mmol) dissolved in DMF (1.5 mL). To this mixture, CuSO4·5H2O (10 mol % per alkyne, in 0.1 mL of deionized water) was added, followed by the addition of sodium ascorbate (15 mol % per alkyne) after 2 min. The reaction was stirred at 40 °C for 10 h. Upon completion, the crude product was purified via dialysis using a 1 kDa MWCO membrane against deionized water for 12 h. The resulting aqueous solution was lyophilized to yield compound 7 as a white solid in an 88% yield.
^1^H NMR (500 MHz, DMSO-d6) δ 8.71 (s, 10H, Drug-Ar H), 8.26–7.66 (m, 177H, Dendrimer-internal amide H and Drug-Ar H), 7.10 (s, 10H, Drug-Ar H), 5.53 (m, 20H, Drug-CH2), 5.37–5.23 (m, 10H, Drug-CH), 4.77 (bs, Dendrimer-OH), 4.28 (m, 20H, Drug-CH2), 4.08–3.95 (m, 26H, Dendrimer-Ester CH2), 3.56–3.42 (m, 156H, Dendrimer-CH2), 3.14 (m, 248H, Dendrimer-CH2), 2.69 (m, 248H, Dendrimer-CH2), 2.47 (m, 126H, Dendrimer-CH2), 2.24 (m, 248H), 2.02–1.70 (m, 34H, Drug-CH2), 1.66–1.42 (m, 40H, Linker-CH2 and Drug-CH2), 1.39–1.14 (m, 42H, Linker-CH2), 1.05–0.85 (m, 30H, Drug-CH3) (Figure S7).
HPLC 98.57%, retention 15.632 min (Figure S8).
PD-Campto (7) (20.0 mg, 1.0 equiv, 0.001 mmol) was dissolved in DI water (150 μL), followed by the addition of DBCO-C6-CTT1298 (8) (2.5 mg, 2.5 equiv, 0.002 mmol). The reaction mixture was stirred at room temperature for 4 h. Reaction progress was tracked with HPLC. Upon completion, the crude product was purified by dialysis against deionized water using a 1 kDa molecular weight cutoff membrane for 12 h. The aqueous solution was lyophilized to afford PD-Campto-CTT1298 (9) in a 93% yield.
^1^H NMR (500 MHz, DMSO-d6) δ 8.71 (s, 10H, Drug-Ar H), 8.09–7.72 (m, 198H, Dendrimer-internal amide H, Drug-Ar H and Ligand-Ar H), 7.14–7.08 (s, 10H, Drug-Ar H), 5.53 (m, 20H, Drug-CH2), 5.32 (m, 20H, Drug-CH), 4.78 (bs, Dendrimer-OH), 4.28 (m, 29H, Drug-CH2), 4.01 (m, 40H, Dendrimer-Ester CH2), 3.42 (m, 275H, Dendrimer-CH2), 3.18–3.08 (m, 251H, Dendrimer-CH2), 2.72–2.65 (m, 225H, Dendrimer-CH2), 2.46 (m, 131H, Dendrimer-CH2), 2.24 (m, 248H, Dendrimer-CH2), 1.98–1.71 (m, 52H, Drug-CH2 and Ligand-CH2), 1.53 (s, 41H, Linker-CH2, Drug-CH2 and Ligand-CH2), 1.36–1.13 (m, 48H, Linker-CH2, Drug-CH2 and Ligand-CH2), 1.13–0.74 (m, 27H, Drug-CH3) (Figure S9).
^31^P NMR (202 MHz, DMSO-d6) δ 6.85 (Figure S10).
HPLC 99.55%, retention 15.420 min (Figure S11).
Size: 13.9 ± 1.99 nm (Figure S12); Zeta −4.72 ± 0.43 mV (Figure S13).
Azido Hexanoic Acid (5) (165 mg, 15.0 equiv, 1.05 mmol) was dissolved in anhydrous DMF (5 mL), followed by the addition of EDC· HCl (377 mg, 28.0 equiv, 1.96 mmol). The mixture was stirred for 15 min at room temperature. This activated solution was then added dropwise to a separately prepared solution of PD (4) (1.0 g, 1.0 equiv, 0.07 mmol) in anhydrous DMF (5 mL) under continuous stirring. Subsequently, DMAP (54 mg, 7 equiv, 0.44 mmol) was added, and the reaction mixture was stirred at room temperature for 24 h. The reaction completion was confirmed by the shift in the retention time of the chromatogram in HPLC. The crude product was purified by dialysis against DI water using a 1 kDa MWCO dialysis membrane for 24 h. The aqueous solution was lyophilized to afford PD-azide-B (10) in a 79% yield.
^1^H NMR (500 MHz, DMSO-d6) δ 8.08–7.77 (m, 125H, Dendrimer-internal amide H), 4.80–4.63 (m, Dendrimer-OH), 4.07–3.92 (m, 26H, Dendrimer-Ester CH2), 3.55–3.24 (m, 209H, Dendrimer-CH2), 3.21–2.96 (m, 237H, Dendrimer-CH2), 2.92–2.72 (m, 52H, Linker-CH2), 2.72–2.54 (m, 251H, Dendrimer-CH2), 2.54–2.34 (m, 159H), 2.32–2.27 (m, 33H, Dendrimer-CH2), 2.26–2.07 (m, 248H, Dendrimer-CH2), 1.66–1.45 (m, 56H, Linker-CH2), 1.38–1.22 (m, 28H, Linker-CH2) (Figure S14).
HPLC 98.44%, retention 14.410 min (Figure S15).
To a stirred solution of compound 10 (300 mg, 1.0 equiv, 0.018 mmol) in DMF (1.5 mL) was added a solution of compound 3 (83 mg, 10.5 equiv, 0.19 mmol) dissolved in DMF (1.5 mL), followed by the addition of CuSO4·5H2O (10 mol % per acetylene) dissolved in DI water (0.1 mL). After 2 min, sodium ascorbate (15 mol % per acetylene) was added, and the reaction was stirred at 40 °C for 10 h. Upon completion, the crude reaction mixture was purified by dialysis against DI water using a 1 kDa MWCO dialysis membrane for 12 h. The aqueous solution was lyophilized to afford PD-Campto′ 11 in 85% yield.
^1^H NMR (500 MHz, DMSO-d6
δ 8.71 (s, 10H, Drug-Ar H), 8.19–7.70 (m, 173H, Dendrimer-internal amide H and Drug-Ar H), 7.07 (s, 10H, Drug-Ar H), 5.53 (m, 14H, Drug-CH2), 5.32 (m, 10H, Drug-CH), 4.76 (bs, Dendrimer-OH), 4.28 (m, 20H, Drug-CH2), 4.01 (m, 29H, Dendrimer-Ester CH2), 3.43 (m, 108H, Dendrimer-CH2), 3.14 (m, 250H, Dendrimer-CH2), 2.68 (m, 245H, Dendrimer-CH2), 2.46 (m, 101H, Dendrimer-CH2), 2.24 (m, 260H, Dendrimer-CH2), 1.97–1.71 (m, 69H, Drug-CH2), 1.54 (m, 48H, Linker-CH2 and Drug-CH2), 1.37–1.12 (m, 46H, Linker-CH2 and Drug-CH2), 1.02–0.86 (m, 30H, Drug-CH3) (Figure S16).
HPLC 99.32%, retention 16.095 min (Figure S17).
PD-Campto′ (11) (20.0 mg, 1.0 equiv, 0.001 mmol) was dissolved in DI water (150 μL), followed by the addition of DBCO-C6-CTT1298 (8) (2.3 mg, 2.5 equiv, 0.002 mmol). The reaction mixture was stirred for 4 h at room temperature. Reaction progress was tracked with HPLC. Upon completion, the crude product was purified by dialysis against DI water using a 1 kDa MWCO dialysis membrane for 12 h. The aqueous solution was lyophilized to afford PD-Campto-CTT1298′ (12) in 91% yield.
^1^H NMR (500 MHz, DMSO-d6) δ 8.71 (s, 10H, Drug-Ar H), 8.46–7.63 (m, 196H, Dendrimer-internal amide H, Drug-Ar H and Ligand-Ar H), 7.11 (s, 10H, Drug-Ar H), 5.53 (m, 21H, Drug-CH2), 5.32 (m, 10H, Drug-CH), 4.80 (bs, Dendrimer-OH), 4.27 (m, 40H, Drug-CH2), 4.01 (m, 48H, Dendrimer-Ester CH2), 3.76–3.59 (m, 105H, Dendrimer-CH2), 3.38–2.95 (m, 250H, Dendrimer-CH2), 2.67 (m, 247H, Dendrimer-CH2), 2.46 (s, 102H, Dendrimer-CH2), 2.23 (m, 248H, Dendrimer-CH2), 2.00–1.70 (m, 48H, Drug-CH2 and Ligand-CH2), 1.53 (s, 34H, Linker-CH2, Drug-CH2 and Ligand-CH2), 1.40–1.14 (m, 46H, Linker-CH2, Drug-CH2 and Ligand-CH2), 1.09–0.85 (m, 30H, Drug-CH3) (Figure S18).
^31^P NMR (202 MHz, DMSO-d6) δ 6.85 (Figure S19).
HPLC 99.62%, retention 15.337 min (Figure S20).
PD-Campto-CTT1298′ (12) (20.0 mg, 1.0 equiv, 0.000 mmol) was dissolved in DI water (150 μL) and stirred at room temperature. To this solution, Cy5-DBCO (13) (2.3 mg, 2.5 equiv, 0.002 mmol) was added, and the reaction mixture was allowed to stir for 48 h at room temperature. The completion of the reaction was tracked using HPLC. The crude product was purified by dialysis against deionized water using a 1 kDa molecular weight cutoff (MWCO) membrane for 12 h. The aqueous solution was lyophilized to afford compound PD-Campto-CTT1298-Cy5 (14) in 81% yield.
^1^H NMR (500 MHz, DMSO-d6) δ 8.70 (s, 10Ḩ Drug-Ar H), 8.45–8.36 (m, 13H, Cy5-Ar H), 8.12–7.67 (m, 236H, Dendrimer-internal amide H, Drug-Ar H, Ligand-Ar H and Cy5-Ar H), 7.37–7.34 (m, 10H, Cy5-Ar H), 7.11 (s, 20H, Drug-Ar H), 6.64–6.59 (m, 5H, Cy5-Ar H), 6.36–6.33 (m, 5H, Cy5-Ar H), 5.61–5.48 (m, 20H, Drug-CH2), 5.32 (m, 11H, Drug-CH2), 4.79 (bs, Dendrimer–OH), 4.27 (m, 51H, Drug-CH2), 4.20–4.10 (m, 48H, Cy5-CH2), 4.06–3.95 (m, 63H, Dendrimer-Ester CH2), 3.18–3.07 (m, 251H, Dendrimer-CH2), 2.68 (s, 266H, Dendrimer-CH2), 2.46 (s, 134H, Dendrimer-CH2), 2.23 (s, 248H, Dendrimer-CH2), 2.08–2.02 (m, 16H, Ligand-CH2 and Cy5-CH2), 1.94–1.88 (m, 17H, Ligand-CH2 and Cy5-CH2), 1.83–1.47 (m, 166H, Drug-CH2, Ligand-CH2, and Cy5-CH2), 1.41–1.15 (m, 93H, Drug-CH2, Linker-CH2, Ligand-CH2, and Cy5-CH2), 1.11–0.89 (m, 86H Drug-CH3 and Cy5-CH2) (Figure S21).
HPLC 99.89%, retention 18.121 min (Figure S22).
Drug release studies were conducted in vitro under both plasma conditions (100% human plasma) and intracellular conditions (citrate buffer pH 5.5 with porcine liver esterase). PD-Campto-CTT1298 was dissolved at a concentration of 1 mg/mL in each buffer. For the intracellular condition, 3 mg of solid PLE (specific 16 units/mg) was dissolved in 3 mL of citrate buffer to achieve an enzyme concentration of 16 U/mL. All solutions were incubated at 37 °C with continuous shaking to simulate physiological conditions. The esterase solution was replenished every 3 days to maintain consistent enzymatic activity over the 15-day study period. At designated time intervals, samples were withdrawn, promptly quenched with an equal volume of methanol, and stored at −20 °C until further use. The released drug was subsequently analyzed using HPLC, and the extent of drug release was determined by comparison to the standard curve established for free Campto on HPLC.
PD-Campto-CTT1298 was formulated at 1 mg/mL in PBS and sterile-filtered through 0.4 μm filters. The formulations were kept at 40 °C, room temperature, and 4 °C. The aliquots were analyzed via HPLC for purity at 1, 4, 7, and 14-day time points.
For in vitro evaluation of PD-Campto-CTT1298, the human PC cell lines PC3-PIP and C4–2B were used due to their relevance in modeling PSMA-expressing tumors. PC3-PIP, a derivative of the PSMA-negative PC3 line, is genetically engineered to stably overexpress prostate-specific membrane antigen (PSMA), thereby serving as an ideal model for testing PSMA-targeted drug delivery systems.^35,36^ The C4–2B cell line on the other hand, derived from LNCaP, is androgen receptor–positive and represents a castration-resistant PC phenotype with clinically relevant PSMA expression levels.^37,38^ The use of both lines provides a strong platform to evaluate PSMA-mediated uptake, internalization, and cytotoxic efficacy of therapeutic agents at distinct stages of PC progression. PC3-PIP and C4–2B cells were cultured in RPMI-1640 medium (VWR) supplemented with 10% fetal bovine serum (FBS; VWR) and 1% penicillin-streptomycin (VWR), at 37 °C and 5% CO2. Cells were passaged at 70–80% confluence using 0.05% trypsin-EDTA (VWR) and subcultured at a 5 split ratio in T-75 flasks (VWR). Both cell lines were regularly tested for mycoplasma contamination and maintained within passage numbers 5–25 to ensure experimental reproducibility. PSMA expression in PC3-PIP and C4–2B has been previously validated by our group using Western blotting and was consistent with their established use as PSMA-overexpressing (PC3-PIP) and clinically relevant, lower-PSMA (C4–2B) prostate cancer models.^38,39^
Uptake of PD-Campto-CTT1298-Cy5 was investigated to evaluate receptor-mediated internalization. For flow cytometry, 2.5 × 10^5^ cells per sample (n = 3) were incubated with PD-Campto-CTT1298-Cy5 (25 μg/mL) for 1, 12, and 24 h at 37 °C. For PSMA blocking, cells were preincubated with 500 nM CTT1057^40^ (an irreversible PSMA inhibitor) for 1 h before treatment with PD-Campto-CTT1298-Cy5 for 6 h. At each time point, cells were collected by centrifugation (1200 rpm, 5 min), washed 3× with ice-cold PBS to remove unbound conjugate, and fixed with 4% paraformaldehyde. Samples were resuspended in FACS buffer and analyzed on an Attune NxT Flow Cytometer using a 640 nm excitation laser and a 670/14 nm emission filter for Cy5 detection. Gating was established using unstained controls, and doublets were excluded based on FSC/SSC parameters. All experiments were conducted in biological triplicate.
To investigate PSMA-specific uptake and internalization pathways of PD-Campto-CTT1298-Cy5, PC3-PIP and C4–2B cells were pretreated with pharmacological inhibitors prior to confocal imaging following our previous reports.^23^ For PSMA blocking, cells were preincubated with 5 μM CTT1057^41^ (an irreversible PSMA inhibitor) for 1 h before treatment with PD-Campto-CTT1298-Cy5 (50 μg/mL) for 6 h at 37 °C. To assess the major endocytic pathways, cells were pretreated for 30–60 min with one of the chlorpromazine (CPZ, 10 μg/mL; ~30 μM) to inhibit clathrin-mediated endocytosis, and methyl-β-cyclodextrin (MβCD, 2.5 mg/mL; ~2 mM) to disrupt caveolae/lipid raft-mediated uptake. To visualize subcellular localization, cells were costained with LysoTracker Green DND-26 (Thermo Fisher, 75 nM) during the final 30 min of PD-Campto-CTT1298-Cy5 treatment. After incubation, all slides were washed with ice-cold PBS, fixed with 4% paraformaldehyde, and mounted with VECTASHIELD Antifade Mounting Medium with DAPI. Imaging was performed using a Leica SP5 confocal microscope with filter sets for DAPI (Ex: 358 nm; Em: 461 nm), LysoTracker Green (Ex: 504 nm; Em: 511 nm), and Cy5 (Ex: 649 nm; Em: 670 nm).
To assess the cytotoxic potential of PD-Campto-CTT1298 and compare it with the free drug, Campto, cells were seeded in 96-well plates at a density of 25,000 cells per well and incubated overnight to allow for adherence. The following day, cells were treated with various concentrations of free Campto, PD-Campto-CTT1298, and DMSO vehicle control or left untreated. After 48 h of incubation at 37 °C, cell viability was determined using the CellTiter-Glo Luminescent Cell Viability Assay (Promega) according to the manufacturer’s instructions. Luminescence, which reflects intracellular ATP content as an indicator of metabolically active cells, was measured by using a FLUOstar Omega microplate reader. Data were expressed as mean ± standard deviation (SD) from at least three independent experiments. Statistical comparisons between free Campto and PD-Campto-CTT1298 were calculated using a two-tailed, unpaired Student’s t-test with unequal variances.
To evaluate the effects of PD-Campto-CTT1298 on cellular proliferation, PC3-PIP and C4–2B cells were seeded into 96-well plates at a density of 25,000 cells per well and allowed to adhere overnight. The next day, cells were treated with varying concentrations of free Campto, PD-Campto-CTT1298, DMSO control, or left untreated for a total of 48 h at 37 °C. During the final 24 h of treatment, BrdU (5-bromo-2′-deoxyuridine) was added to the culture medium to label newly synthesized DNA. Cell proliferation was then quantified using the BrdU Cell Proliferation ELISA Kit (chemiluminescent; Abcam), following the manufacturer’s protocol. Briefly, cells were fixed, DNA was denatured, and BrdU incorporation was detected using a monoclonal anti-BrdU antibody, followed by a horseradish peroxidase-conjugated secondary antibody. After substrate addition, chemiluminescence was measured by using the FLUOstar Omega microplate reader. Proliferation was expressed as a fold change relative to the untreated control. Results represent the mean ± SD from the experiment performed in triplicate. Statistical significance was assessed using a two-tailed t-test with unequal variances.
To investigate the apoptosis-inducing potential of PD-Campto-CTT1298 in PC cells, a dual-staining flow cytometric approach was used with Annexin V-FITC and propidium iodide (PI). PC3-PIP and C4–2B cells were seeded at a density of 1 × 10^6^ cells per well in 6-well plates and allowed to adhere overnight. The following day, cells were treated with increasing concentrations (50, 100, and 250 μg/mL) of free Campto or PD-Campto-CTT1298 for 24 h at 37 °C. Hydrogen peroxide (H2O2, 500 μM and 2 mM) served as a positive control for apoptosis and necrosis. Following treatment, cells were collected, washed 2× with ice-cold PBS, and resuspended in 100 μL of 1× binding buffer provided in the Annexin V Apoptosis Detection Kit (SouthernBiotech, FITC). Samples were stained with 5 μL of Annexin V-FITC and 1 μL of PI (100 μg/mL stock) and incubated in the dark at room temperature for 30 min. Prior to acquisition, 400 μL of additional binding buffer was added to each sample and mixed gently. Flow cytometric analysis was performed on an Attune NxT Flow Cytometer using a 488 nm laser for excitation. Emission signals were collected at 530/30 nm (Annexin V-FITC, BL-1) and 585/16 nm (PI, YL-1). Compensation controls and unstained samples were used to define gating. Forward and side scatter parameters were used to eliminate debris and cell aggregates. Populations were categorized as viable (Annexin V^−^/PI^−^), early apoptotic (Annexin V^+^/PI^−^), late apoptotic (Annexin V^+^/PI^+^), and necrotic (Annexin V^−^/PI^+^). Data were representative of experiment performed in triplicate. Statistical comparisons were made using an unpaired two-tailed t-test with unequal variances.
Disruption of mitochondrial membrane potential (ΔΨm) in PC cells was assessed by using the JC-1 Mitochondrial Membrane Potential Assay Kit (MedChemExpress). PC3-PIP and C4–2B cells were seeded at a density of 1 × 10^6^ cells per well in 6-well plates and incubated overnight. The following day, cells were exposed to increasing concentrations (50, 100, and 250 μg/mL) of free Campto or PD-Campto-CTT1298 for 24 h at 37 °C. Carbonyl cyanide 3-chlorophenylhydrazone (CCCP, 50 μM) was used as a positive control for mitochondrial depolarization. Following treatment, cells were collected, washed 2× with ice-cold PBS, and resuspended in 1 mL of PBS. JC-1 dye was added to each sample at a final concentration of 2 μM, and the samples were incubated at 37 °C for 1 h in the dark. After being stained, cells were washed and resuspended in 500 μL of PBS for flow cytometric analysis. Fluorescence was detected using an Attune NxT Flow Cytometer equipped with a 488 nm excitation laser. JC-1 monomers (depolarized mitochondria) emitted green fluorescence (BL-1, 530/30 nm), while JC-1 aggregates (polarized mitochondria) emitted red fluorescence (YL-1, 585/16 nm). Unstained and single-stained controls were used to establish compensation settings and gating. The experiment was performed in triplicate, and statistical significance was determined using an unpaired, two-tailed t-test with unequal variances.
Caspase-3/7 activity was measured to assess apoptotic pathway activation following treatment with PD-Campto-CTT1298. PC3-PIP and C4–2B cells were seeded at 1 × 10^6^ cells per well in 6-well plates and incubated overnight. The next day, cells were treated with 50, 100, or 250 μg/mL of free Campto or PD-Campto-CTT1298 for either 12 or 24 h at 37 °C. Hydrogen peroxide (H2O2, 500 μM) was used as a positive control for caspase induction, while untreated cells served as negative controls. After treatment, cells were collected, washed 2× with cold PBS, and resuspended in 100 μL of assay buffer. Caspase-3/7 fluorogenic substrate (Revvity, FITC), 5 μL, was added to each sample, followed by incubation at 37 °C for 1 h. Cells were subsequently washed and resuspended in 500 μL of FACS buffer containing propidium iodide (PI). Fluorescence detection was performed using an Attune NxT Flow Cytometer with excitation at 488 nm. Caspase-3/7 activity was detected in BL-1 (530/30 nm) and PI was detected in YL-1 (585/16 nm). Gating was established using unstained, caspase-only, and PI-only controls. Cells were classified into four viable (Caspase-3/7^−^/PI^−^), early apoptotic (Caspase-3/7^+^/PI^−^), late apoptotic (Caspase-3/7^+^/PI^+^), and necrotic (Caspase-3/7^−^/PI^+^). The experiment was performed in triplicate at each time point, and statistical analysis was calculated using an unpaired, two-tailed t-test with unequal variances.
To evaluate hypoxia induction following treatment with PD-Campto-CTT1298, the expression of the hypoxia-responsive marker carbonic anhydrase IX (CA IX) was assessed by using the IVISense Hypoxia CA IX 680 fluorescent probe. PC3-PIP and C4–2B cells were seeded in 6-well plates at a density of 1 × 10^6^ cells per well and incubated overnight. The following day, cells were treated with free Campto or PD-Campto-CTT1298 at concentrations of 50, 100, or 250 μg/mL for 24 h at 37 °C. Hydrogen peroxide (H2O2, 250 μM) was included as a positive control to validate hypoxia induction. Following treatment, cells were incubated with IVISense Hypoxia CA IX 680 (10 μM final concentration) for 1 h at 37 °C. After staining, cells were washed 2× with ice-cold PBS to remove excess dye and fixed with 4% paraformaldehyde. Samples were resuspended in FACS buffer and analyzed using an Attune NxT Flow Cytometer equipped with a 640 nm excitation laser and a 680/30 nm emission filter for detection of the CA IX 680 signal. Unstained and single-color controls were used for gating. CA IX expression levels were quantified by mean fluorescence intensity (MFI) and reported as fold change relative to untreated controls. The experiment was performed in triplicate and statistical comparisons were performed using an unpaired, two-tailed t-test with unequal variances.
To evaluate the role of reactive oxygen species (ROS) in mediating the cytotoxic effects of PD-Campto-CTT1298, confocal imaging of DCFDA-stained PC cells was performed following our previously published report with minor modifications.^41^ PC3-PIP and C4–2B cells were seeded into chamber slides at 1 × 10^5^ cells per well and allowed to adhere overnight. The next day, cells were incubated with 20 μM DCFDA (Abcam) prepared in phenol red–free RPMI-1640 medium for 1 h at 37 °C to enable probe internalization and deacetylation to the ROS-sensitive form. Following DCFDA loading, cells were washed twice with warm PBS and treated with 250 μg/mL of either free Campto or PD-Campto-CTT1298 for 12 h. A positive control group, 100 μM hydrogen peroxide (H2O2), was used to induce oxidative stress. After treatment, cells were gently washed with PBS and fixed with 4% paraformaldehyde and mounted using a VECTASHIELD Antifade Mounting Medium. Imaging was conducted using a Leica SP5 confocal laser scanning microscope equipped with a FITC filter set (Ex: 488 nm; Em: 529 nm). Green fluorescence intensity, proportional to intracellular ROS accumulation, was captured by using identical settings across all treatment groups.
To determine whether PD-Campto-CTT1298 induces autophagy in PC cells, a fluorescence-based autophagy detection assay was performed using the CYTO-ID Autophagy Detection Kit (Enzo Life Sciences) according to the manufacturer’s instructions. PC3-PIP and C4–2B cells were seeded into 6-well plates at a density of 1 × 10^6^ cells per well and allowed to adhere overnight. The next day, cells were treated with free Campto or PD-Campto-CTT1298 at concentrations of 50, 100, or 250 μg/mL for 24 h at 37 °C. A positive control group was treated with the autophagy inducer rapamycin (1 μM) to validate assay performance. After treatment, cells were harvested by trypsinization, washed twice with assay buffer, and resuspended in 250 μL of fresh buffer containing the CYTO-ID Green Detection Reagent. Samples were incubated at 37 °C for 30 min in the dark, washed, and resuspended in assay buffer for flow cytometric analysis. To assess autophagic flux, a subset of samples was cotreated with chloroquine (25 μM) during the final 4 h of the experiment. Samples were analyzed on an Attune NxT Flow Cytometer equipped with a 488 nm excitation laser and a 530/30 nm emission filter. Fluorescence signal intensity was used to quantify autophagic vacuole accumulation. Unstained and single-stained controls were used for gating and compensation. The experiment was performed in triplicate, and statistical comparisons were made using a two-tailed t-test with unequal variances.
To assess the effects of PD-Campto-CTT1298 and free Campto on angiogenesis, VEGF-A secretion was measured using a Human VEGF-A ELISA Kit (VWR). PC3-PIP and C4–2B cells (1 × 10^6^ per sample) were seeded in 6-well plates and allowed to adhere overnight at 37 °C. The next day, cells were treated with free Campto or PD-Campto-CTT1298 at concentrations of 50, 100, or 250 μg/mL for 24 h at 37 °C, with untreated cells serving as a negative control. After incubation, the cell culture supernatant was collected and centrifuged (10,000 rpm, 15 min) to remove debris. The supernatant was also filtered through a 0.22 μm PES filter to ensure a clean sample for ELISA analysis. To prevent protein degradation, a protease inhibitor cocktail (Sigma-Aldrich) was added to the supernatant at the manufacturer’s recommended concentration. Samples were stored at −80 °C until further analysis. VEGF-A levels were quantified using the Human VEGF-A ELISA Kit (VWR), following the manufacturer’s protocol. Briefly, 100 μL of supernatant was added to precoated 96-well plates, incubated, and processed per kit instructions. Absorbance was measured at 450 nm using a microplate reader, and data were normalized to untreated controls. The experiment was conducted in triplicate, and statistical significance was determined using a two-tailed t-test with unequal variances.
To assess the indirect antiangiogenic effects of PD-Campto-CTT1298 via tumor-secreted factors, a tube formation assay was performed using HUVECs treated with conditioned media derived from drug-treated PC cells. PC3-PIP and C4–2B cells were seeded in 6-well plates at a density of 1 × 10^6^ cells per well and treated with 250 μg/mL of free Campto or PD-Campto-CTT1298 for 24 h at 37 °C. Following treatment, supernatants were collected, centrifuged (10,000 rpm, 15 min) to remove cell debris, and filtered through a 0.22 μm PES syringe filter. The resulting conditioned media (CM) were stored at −80 °C until further use. Separately, a 96-well plate was precoated with 50 μL of extracellular matrix solution provided in the Angiogenesis Assay Kit (Abcam) and incubated at 37 °C for 30 min to allow gel polymerization. HUVECs were seeded at 2 × 10^4^ cells per well in a 1 mixture of complete endothelial growth medium and PC CM (vehicle, Campto, or PD-Campto-CTT1298). Untreated cell supernatant and Suramin (known to disrupt tube formation) were used as controls. Plates were incubated at 37 °C for 24 h, stained using the kit dye for 30 min, and imaged at 5× magnification using a Zeiss Axiovert fluorescence microscope. Images were analyzed using ImageJ (NIH, version 1.54j) with a custom macro tailored for angiogenesis assessment. The plugin automatically quantified key parameters such as tube length, nodes, junctions, master segments, meshes, segments, and branches to evaluate the antiangiogenic effect of the PD-Campto-CTT1298 dendrimer conjugate on HUVEC cells. All experiments were performed in triplicate with the proper controls.
To determine the therapeutic interaction between PD-Campto-CTT1298 and Free Olaparib (Free OP), combination treatment studies were conducted. PC3-PIP and C4–2B cells were seeded at 25,000 cells per well in 96-well plates and incubated overnight to allow adherence. The following day, cells were treated with Free OP or a 1 combination of PD-Campto-CTT1298 and OP at concentrations ranging from 50 to 500 μg/mL. After 48 h at 37 °C, cell viability was measured using the CellTiter-Glo Luminescent Assay, and luminescence data were collected using a microplate reader. Viability was calculated relative to that of the untreated controls. Previously collected data for free Campto and PD-Campto-CTT1298 monotherapies under identical conditions were used for reference to assess combination-specific effects. To characterize the interaction between PD-Campto-CTT1298 and OP, three established synergistic models were applied. The experiment was performed in triplicate with statistical analysis using an unpaired, two-tailed t-test with unequal variances. Synergy scores were computed using different synergy analysis softwares (Table 1).
To synthesize PD-Campto-CTT1298, we first modified the hydroxyl group of Campto (1) at the 20-position by conjugating it with hexynoic acid (2) via Steglich esterification, yielding Campto-Hexyne (3) (Figures 1A and S1). This modification facilitated the subsequent click conjugation of Campto. Previous studies have demonstrated that esterification at the hydroxyl group at the 20-position can act as a prodrug strategy, enabling intracellular release of the active drug.^42^ Ester derivatives of Campto have been shown to enhance pharmacokinetics and improve drug delivery without significantly compromising cytotoxic activity.^43^ The successful conjugation of the hexyne moiety on Campto was confirmed by the appearance of aliphatic protons from the linker between δ1.7 and δ2.3 ppm and the acetylene triplet at δ2.9 ppm in the ^1^H NMR spectrum (Figure 1C, blue spectrum). HPLC analysis showed a shift in retention time from 14.7 min for Campto to 20.6 min for Campto-Hexyne, with a purity exceeding 95% (Figures 2A and S3).
Next, we partially modified the surface hydroxyl groups of PD (4). PD is neutral and exhibits nontoxicity and biocompatibility, making it highly suitable for applications in targeted drug delivery.^28,44^ The modification of PD (4) was achieved by reacting the hydroxyls on its surface with azido-hexanoic acid (5) under Steglich esterification conditions to form PD-Azide (6) (Figure 1A). The reaction resulted in the attachment of 12 azide arms to the surface of the dendrimer. The incorporation of the azide linkers was confirmed by comparing the linker protons (24 and 48H) between δ1.2 and δ1.5 ppm with the internal amide protons (124H) of PD, which appeared between δ7.7 and δ8.0 ppm, using proton integration analysis (Figures 1B and S5). HPLC analysis displayed a purity level of >99% (Figures 2A and S6). Subsequently, we conjugated Campto-Hexyne (3) and PD-azide (6) by employing CuAAC, an orthogonal chemistry technique, to synthesize PD-Campto (7). The shift in retention time of the HPLC chromatogram from 12.5 min (PD-Azide) to 15.6 min and the disappearance of the Campto-Hexyne (3) peak at 20.6 min suggested the completion of the click (Figure 2A). The number of attached Campto molecules was confirmed through the comparative integration of aromatic protons from the drug at δ7.0 and δ8.7 ppm and the aliphatic protons of the dendrimer at δ2.2 ppm (Figures 1B and S7). Finally, PD-Campto (7) was conjugated with the PSMA-targeting ligand (CTT1298) by using the SPAAC click reaction to afford PD-Campto-CTT1298 (9). For this, CTT1298 was first modified by introducing a DBCO ring using a previously published procedure,^31^ yielding DBCO-C6-CTT1298 (8). The strained alkyne on the DBCO ring readily reacts with the azide groups, forming a stable triazole linkage via the SPAAC reaction. SPAAC offers several advantages, including high specificity, bioorthogonality, and the ability to provide precise ligand or drug loading. SPAAC enabled the successful conjugation of CTT1298 onto the dendrimer’s surface without the need for any additional reagents to yield PD-Campto-CTT1298 (9) with ~2 ligand molecules attached. This was confirmed by the comparative integration of aromatic protons from the ligand and drug between δ7.0 and δ8.7 ppm and aliphatic protons (248H) from the dendrimer at δ2.2 ppm (Figures 1B and S9). Furthermore, a signal at δ6.85 ppm in the ^31^P NMR spectrum validated the conjugation of CTT1298 to the dendrimer (Figure S10). The HPLC chromatogram exhibited a shift in retention time, transitioning from ~15.6 min for PD-Campto to ~15.4 min for PD-Campto-CTT1298 upon the conjugation of the PSMA-targeting ligand (Figure 2A). The HPLC purity level of PD-Campto-CTT1298 exceeded 99% (Figures 2A and S11). The size and zeta potential distribution were analyzed by using dynamic light scattering (DLS). The hydrodynamic diameter of PD-Campto-CTT1298 was found to be 13.90 ± 1.99 nm and the zeta potential distribution was −4.72 ± 0.43 mV (Figures 2B, i and ii, S12 and S13). The physicochemical properties of PD-Campto-CTT1298 are presented in Figure 3A, (i). While Campto is practically insoluble at room temperature, its conjugation to the dendrimer significantly improved the water solubility (Figure 3A, (ii)). The aqueous solubility of PD-Campto-CTT1298 was found to be ~180 mg/mL, with ~15 wt % drug loading, which translates to ~27.5 mg/mL on a Campto basis.
We evaluated the shelf stability of the PD-Campto-CTT1298 (1 mg/mL Campto-equivalent concentration) in PBS at 4 °C, room temperature (25 °C), and 40 °C over a 14-day period. The formulation maintained its stability with more than 99% HPLC purities over the experimental period at all temperatures (Figure S23). No changes in retention time were observed throughout the 14 days, indicating the stability of the formulation under these conditions.
We also evaluated the in vitro drug release profile of the conjugate under conditions that mimic both the extracellular environment (PBS buffer and 100% human plasma) and the intertumoral microenvironment (pH 5.5 citrate buffer and PLE) (Figure 3A (iii) and (iv)). Campto is conjugated to the dendrimer via an ester bond at its hydroxyl group at position 20, enabling pH- and esterase-responsive release. This strategic design ensures controlled drug release within the tumor and intracellular compartments while minimizing systemic exposure beyond the prostate tumor site. In PBS buffer, we observed negligible drug release. To simulate in vivo conditions, we further evaluated the stability of PD-Campto-CTT1298 in human plasma and found that less than 20% of the drug was released over 48 h. Under intracellular conditions, the conjugate exhibited a sustained and gradual release profile, with ~15% of the drug released in the first 8 h, ~25% at 48 h, and approximately 85% over 15 days (Figure 3A (iii)). The observed intracellular release enhances the therapeutic potential of PD-Campto-CTT1298 for targeted PC treatment.
A major challenge in the clinical translation of nanoparticle-based therapeutics is the lack of reproducibility and scalability of their synthesis. To address these limitations, we developed an efficient and accelerated synthetic approach for PD-Campto-CTT1298, allowing the precise characterization of intermediates at each step, as demonstrated above. To further evaluate synthetic reproducibility, we prepared three 200 mg-scale batches and analyzed their consistency using ^1^H NMR and HPLC. The ^1^H NMR spectra of the three independent batches (Figure 3B, left) demonstrated reproducible synthesis, and drug and ligand loading. Additionally, the HPLC chromatograms of these batches displayed peaks at the same retention time (~15.6 ± 0.5 min), and purity >98%, confirming batch-to-batch consistency (Figure 3B, right).
To investigate the in vitro uptake via confocal microscopy and fluorescence spectroscopy, we further attached Cy5 on the surface of PD-Campto-CTT1298 (Figure 4A). Hydroxyl groups on the surface of PD (4) were partially modified by reacting with azido-hexanoic acid (5) in the presence of EDC-DMAP to obtain the azide-bearing dendrimer PD-Azide′ (10) with 14 azide arms. The number of azide linkers was confirmed by comparing the linker protons (28H and 56H) between δ1.3 and δ1.6 ppm with the internal amide protons (124H) of PD, which appeared between δ7.8 and δ8.1 ppm, using proton integration analysis (Figure S14). Next, Campto-Hexyne (3) was conjugated with PD-Azide′ (10) via a CuAAC reaction to form PD-Campto′ (11). The product formation was verified using the comparative proton integration method and the observed shift in retention time in HPLC (Figures S16 and S17). Subsequently, DBCO-C6-CTT1298 (8) (~2 molecules) was coupled to the dendrimer’s surface using SPAAC chemistry. This reaction yielded PD-Campto-CTT1298′ with 2 free azides left on the surface. The PD-Campto-CTT1298′ (12) was conjugated with DBCO-Cy5 (13) via the SPAAC reaction, yielding the fluorescently labeled PD-Campto-CTT1298-Cy5 (14). The successful attachment of Cy5 was confirmed through the identification and comparative integration of Cy5-specific protons between δ6.3 and δ7.5 ppm, along with aromatic and amide protons from the dendrimer-drug conjugate between δ7.6 and δ8.1 ppm in the ^1^H NMR spectrum (Figures 4B (i) and S21). Proton integration analysis indicated two Cy5 molecules coupled on the dendrimer. The HPLC chromatogram displayed a shift in retention time from 15.3 to 18.1 min upon Cy5 conjugation with >99% purity (Figures 4B (ii) and S22).
To assess the uptake kinetics and specificity of PD-Campto-CTT1298, a time-dependent uptake study using the Cy5-labeled conjugate was performed in the PC3-PIP and C4–2B PC cell lines. PSMA expression is considerably lower in C4–2B cells compared to PC3-PIP cells, as confirmed by previous studies.^36^ Flow cytometry analysis confirmed a time-dependent increase in mean fluorescence intensity (MFI), consistent with rapid internalization and sustained intracellular retention over 24 h (Figure 5). In PC3-PIP cells (Figures 5A,C and S24), PD-Campto-CTT1298-Cy5 showed rapid uptake within 1 h (MFI: 1803 ± 125, p < 0.01 vs untreated), indicating efficient PSMA binding and early internalization. Uptake increased significantly by 12 h (MFI: ~25452 ± 579, p < 0.001 vs 1 h) and continued to rise by 24 h (MFI: ~39285 ± 2469, p < 0.01 vs 12 h), suggesting sustained intracellular accumulation, potentially reflecting endosomal or lysosomal retention. In C4–2B cells (Figures 5A,C and S25), a similar time-dependent trend was observed, though overall uptake was lower. MFI increased from ~684 ± 180 at 1 h to ~12977 ± 1639 at 12 h and ~20317 ± 1766 at 24 h (p < 0.01 across time points). Despite the increase, the fluorescence intensity remained significantly lower than that observed in PC3-PIP cells at all time points. To confirm PSMA specificity, a blocking study was performed using CTT1057, an irreversible PSMA inhibitor with nanomolar affinity (Figure 5B,D). In PC3-PIP cells, CTT1057 pretreatment significantly reduced uptake at 6 h (MFI: ~998 ± 145, p < 0.05 vs unblocked PD-Campto-CTT1298-Cy5), validating PSMA-mediated targeting as the dominant internalization route. However, uptake remained significantly higher than in untreated controls (p < 0.01), indicating residual nonspecific uptake, possibly due to nonreceptor-mediated endocytosis or enhanced permeability and retention (EPR) effects, both of which are well-established features of dendrimer-based nano-therapeutics.^45,46^ In contrast, C4–2B cells showed a drastic decrease in uptake with CTT1057. MFI dropped from ~5551 ± 480 to ~511 ± 196 at 6 h (p < 0.001), effectively eliminating the difference from untreated cells (MFI: ~216 ± 76, ns), further confirming the dependence of uptake on PSMA-mediated targeting mechanisms. Overall, these results confirm that PD-Campto-CTT1298-Cy5 exhibits rapid, sustained, and PSMA-specific uptake with potential advantages conferred by both active targeting and passive accumulation via EPR.
To investigate the cellular uptake mechanism of PD-Campto-CTT1298 in PC3-PIP and C4–2B cells, confocal microscopy was performed using PD-Campto-CTT1298-Cy5 (Figure 5E,F). Confocal imaging demonstrated minimal and nonsignificant internalization in both PC3-PIP and C4–2B cells upon treatment with the inhibitor CTT1057, suggesting that this inhibitor effectively impeded the internalization process. This observation indicates that the uptake of the conjugate predominantly occurs through the receptor-mediated pathway that CTT1057 specifically is blocking. Similarly, complete inhibition of the conjugate internalization was observed when cells were pretreated with CPZ, which strongly supports that PD-Campto-CTT1298 uptake primarily occurs via clathrin-dependent endocytosis. Conversely, pretreatment with MβCD did not result in any reduction of the Cy5 fluorescence signal, indicating that the internalization of PD-Campto-CTT1298 does not involve caveolae-dependent pathways. These findings collectively delineate the cellular uptake route of PD-Campto as primarily mediated through clathrin-coated vesicle pathways, excluding a significant role for caveolae-mediated endocytosis.
To further explore the lysosomal internalization and clearance of the conjugate, confocal microscopy studies were conducted using the Cy5-labeled conjugate in PC3-PIP and C4–2B cells (Figure 5G). Confocal imaging revealed high colocalization of Cy5 fluorescence with LysoTracker green, confirming efficient uptake and subsequent trafficking of PD-Campto-CTT1298 into lysosomes in both cell types. These observations clearly suggest that the PD-Campto-CTT1298 conjugate internalizes predominantly through endocytic pathways, ultimately accumulating within lysosomal compartments.
The efficacy of PD-Campto-CTT1298 was assessed in PC3-PIP and C4–2B PC cell lines using cell viability and BrdU proliferation assays. As shown in Figure 6A,B, PD-Campto-CTT1298 significantly reduced cell viability in a concentration-dependent manner, while free Campto produced only modest effects across the same range. PD-Campto-CTT1298 lowered cell viability across all concentrations especially at higher doses (PC3-PIP: ~48%, p < 0.001; C4–2B: ~55%, p < 0.01 at 500 μg/mL) whereas free Campto-treated cells remained above 75% viability at all the doses, thereby confirming enhanced cytotoxic performance by the dendrimer conjugate. The diminished activity of free Campto is consistent with its well-documented pharmacological limitations. Under physiological conditions, Campto undergoes rapid, pH-dependent hydrolysis of its active lactone ring to an inactive carboxylate form, compromising therapeutic efficacy and drug stability.^47,48^ Additionally, Campto’s poor aqueous solubility promotes precipitation at higher concentrations, limiting bioavailability and cellular uptake.^14^ These drawbacks likely account for the minimal cytotoxicity observed in both PC models. To further confirm that the observed effects arise from the Campto payload, we also tested the empty scaffold, PD-CTT1298, at identical concentrations. PD-CTT1298 alone showed no significant effect on cell viability at any dose, confirming that the dendrimer platform itself is noncytotoxic in both cell lines. In parallel, BrdU incorporation assays supported these findings (Figure 6C,D). PD-Campto-CTT1298 induced a strong and dose-dependent suppression of proliferation, decreasing DNA synthesis by approximately ~70% in PC3-PIP and ~75% in C4–2B cells at 500 μg/mL. Free Campto on the other hand showed only marginal inhibition, particularly at lower doses and failed to achieve comparable effects at higher concentrations. Notably, the antiproliferative effects of PD-Campto-CTT1298 were significantly greater than those of free Campto across all the concentrations tested. These results demonstrate that PD-Campto-CTT1298 significantly outperforms free Campto in both cytotoxicity and antiproliferative activity, likely due to improved drug stability and effective cellular uptake of the dendrimer conjugate. It is important to note that concentrations of PD-Campto-CTT1298 are reported as total conjugate weight (μg/mL), which contains ~15 wt % Campto. Thus, for example, 250 μg/mL of conjugate corresponds to ~37.5 μg/mL Campto equivalent. Despite this lower effective drug content, PD-Campto-CTT1298 consistently produced greater reductions in viability and proliferation compared with free Campto at the same nominal concentrations. This underscores the advantage of the dendrimer formulation in enhancing intracellular delivery and therapeutic efficacy, even at reduced Campto-equivalent doses. Importantly, complementary toxicity studies in nonmalignant HEK-293 cells showed no loss of viability with PD-CTT1298 or PD-Campto-CTT1298 (Figure S26), confirming the biocompatibility of the scaffold and the absence of off-target toxicity.
To evaluate whether PD-Campto-CTT1298 promotes apoptotic cell death in PC, Annexin V-FITC and Propidium Iodide (PI) staining were performed in PC3-PIP and C4–2B cells after 24-h treatment with free Campto or PD-Campto-CTT1298 across a dose range of 50–250 μg/mL. Flow cytometry analysis revealed a clear, dose-dependent increase in both early and late apoptotic populations with PD-Campto-CTT1298 compared to free Campto. In PC3-PIP cells (Figures 7A and S27), treatment with 250 μg/mL PD-Campto-CTT1298 resulted in ~21.7% early and ~41.7% late apoptotic cells, significantly higher than ~9.2% early and 3.0% late apoptosis observed with free Campto (p < 0.01 for both vs free Campto). In C4–2B cells (Figures 7B and S28), PD-Campto-CTT1298 induced ~43.2% early and ~22.6% late apoptosis, compared to ~8.0% and ~1.4% with free Campto at the same concentration (p < 0.001 for both vs free Campto). These apoptotic effects were consistently observed across all tested concentrations, with PD-Campto-CTT1298 showing significantly greater apoptotic induction even at 50 μg/mL (p < 0.05) in C4–2B cells. Correspondingly, PD-Campto-CTT1298 induced a sharp reduction in viable (Annexin V^−^/PI^−^) cell populations at all of the concentrations tested in both cell lines. In contrast, free Campto demonstrated only modest reductions in viability and less consistent apoptotic effects across doses, primarily due to solubility issues. These findings confirm that PD-Campto-CTT1298 induces apoptosis more efficiently than free Campto through improved solubility, intracellular retention, and targeted PSMA-mediated delivery.
To investigate whether mitochondrial membrane depolarization contributes to PD-Campto-CTT1298-induced apoptosis, we used the JC-1 dye assay to monitor ΔΨm in PC3-PIP and C4–2B PC cells. Flow cytometry analysis showed a dose-dependent shift from red (JC-1 aggregates) to green fluorescence (JC-1 monomers), indicating mitochondrial depolarization in both cell lines following 24 h of treatment with PD-Campto-CTT1298 (Figure 7C–F). In PC3-PIP cells, treatment with 250 μg/mL PD-Campto-CTT1298 resulted in a ~80.6% JC-1 monomer population, compared to ~26.2% with free Campto (p < 0.001 vs free Campto; Figure 7D). Similarly, in C4–2B cells, PD-Campto-CTT1298 induced ~57.5% JC-1 monomers, while free Campto yielded ~10.3% (p < 0.001; Figure 7F). This trend continued at lower concentrations and even at 50 μg/mL in the C4–2B cells. The results clearly indicate that PD-Campto-CTT1298 significantly disrupts the mitochondrial membrane potential in a dose-dependent manner, consistent with early intrinsic apoptotic signaling. The statistically significant increase in mitochondrial depolarization across the concentrations tested reinforces the role of ΔΨm collapse as an early event in PD-Campto-CTT1298-induced apoptosis. Unlike free Campto, which suffers from poor solubility and rapid deactivation, PD-Campto-CTT1298 provides stable, water-soluble delivery with enhanced retention and PSMA-mediated uptake. This likely allows for sustained topoisomerase I inhibition and accumulation of DNA damage, culminating in mitochondrial depolarization. These findings align with previous reports that Campto analogs can initiate apoptosis through mitochondrial stress, in part by triggering reactive oxygen species (ROS) and loss of ΔΨm, and eventually leading to downstream caspase activation.^49^ Moreover, enhanced ROS production and mitochondrial collapse have also been implicated in synergistic apoptotic pathways when Campto is combined with other agents.^50^
To investigate the kinetics of apoptosis induction by PD-Campto-CTT1298, we assessed Caspase-3/7 activation in PC3-PIP and C4–2B cells at 12 and 24 h following treatment with 50, 100, and 250 μg/mL of PD-Campto-CTT1298 or free Campto. Flow cytometric analysis was used to gate populations as viable (C3/7^−^/PI^−^), early apoptotic (C3/7^+^/PI^−^), late apoptotic (C3/7^+^/PI^+^), or necrotic (C3/7^−^/PI^+^). In PC3-PIP cells (Figure S29), a modest increase in early apoptotic cells was observed at 12 h, with PD-Campto-CTT1298 showing a clear dose-dependent trend not seen with free Campto. By 24 h, PD-Campto-CTT1298 elicited a significant shift toward late apoptosis, particularly at 250 μg/mL, where ~47% of cells were classified as late apoptotic, compared to only ~13% with free Campto (p < 0.001). Early apoptosis was also elevated across all PD-Campto-CTT1298 doses at 24 h, supporting the presence of sustained apoptotic signaling. The shift from early to late apoptosis over time was statistically significant (p < 0.01 at 50 μg/mL; p < 0.05 at 100 μg/mL), indicating a time-dependent progression of apoptosis. Similarly, in C4–2B cells (Figure S30), PD-Campto-CTT1298 produced an increase in early apoptotic populations by 12 h at all doses tested. At 24 h, there was a significant conversion to late apoptosis, with ~29% late apoptotic cells at 250 μg/mL compared to ~4% in free Campto-treated cells (p < 0.05). Notably, viability declined in both cell lines with increasing concentrations and longer exposure to PD-Campto-CTT1298, highlighting its sustained pro-apoptotic effect. While free Campto showed minor caspase activation at 12 h, the response plateaued at 24 h with minimal progression to late apoptosis, likely due to its poor solubility and reduced intracellular availability. Recent studies confirm that PSMA-targeted Campto delivery either through nanoparticles or small molecule conjugates induces strong, time- and dose-dependent caspase-3 activation, leading to enhanced apoptosis compared to free Campto. These effects are attributed to improved intracellular accumulation and bioavailability via PSMA targeting.^51,52^ Overall, these results demonstrate that PD-Campto-CTT1298 drives a time- and concentration-dependent apoptotic program in both PSMA-positive and androgen-independent PC models. The increase in Caspase-3/7 activation over time is consistent with our Annexin V/PI and mitochondrial depolarization data. Together, these findings support a model in which PSMA-targeted delivery of Campto enhances intracellular retention and sustained apoptotic signaling, resulting in significant therapeutic efficacy compared with free Campto.
To assess the effect of PD-Campto-CTT1298 on hypoxia, PC3-PIP and C4–2B PC cells were treated with free Campto or PD-Campto-CTT1298 at 50, 100, or 250 μg/mL for 24 h and stained with the IVISense Hypoxia CA IX 680 probe. In PC3-PIP cells (Figures 8A and S31), free Campto induced a ~5.8-fold increase in CA IX-associated fluorescence at 50 μg/mL (~2900 MFI; p < 0.05 vs untreated), but this effect was reduced and not significant at 100 μg/mL and 250 μg/mL. In contrast, PD-Campto-CTT1298 treatment resulted in a significant dose-dependent increase in hypoxia marker ~8.0-fold at 50 μg/mL (~4017 MFI; p < 0.05 vs untreated), ~18.6-fold at 100 μg/mL (~9379 MFI; p < 0.01 vs untreated), and ~25.4-fold at 250 μg/mL (~12,758 MFI; p < 0.01 vs untreated). The hypoxia signal induced by PD-Campto-CTT1298 was significantly greater than that induced by free Campto at higher concentrations (p < 0.01 vs free Campto). The same trend was observed in C4–2B cells (Figures 8B and S32). Free Campto resulted in only a ~2.4-fold increase in CA IX signal at 50 μg/mL (~4479 MFI; p < 0.05 vs untreated), but no significant increase at 100 μg/mL or 250 μg/mL. PD-Campto-CTT1298 induced a dose-dependent hypoxic ~3.5-fold at 50 μg/mL (~6559 MFI; p < 0.01 vs untreated), 6.2-fold at 100 μg/mL (, used as a positive control, induced a ~42.8-fold increase in PC3-PIP cells (~21,549 MFI; p < 0.01 vs untreated) and a ~27.6-fold increase in C4–2B cells (~51376 MFI; p < 0.001 vs untreated), validating these experiments. Taken together, these results demonstrated that PD-Campto-CTT1298 induced a significantly stronger and more sustained hypoxic response compared to that of free Campto. This enhanced hypoxia likely results from increased intracellular stress and altered mitochondrial homeostasis, as supported by JC-1 assays showing mitochondrial membrane depolarization and increased apoptosis via Annexin V/PI and caspase-3/7 activation. Recent studies have shown that Campto and its analogs can induce mitochondrial dysfunction and promote hypoxia through reactive oxygen species (ROS) accumulation ultimately sensitizing tumor cells to apoptosis under stress conditions.^53^11628 MFI; p < 0.001 vs untreated), and 222.4-fold at 250 μg/mL (O41,938 MFI; p < 0.001 vs untreated). At each dose, PD-Campto-CTT1298 significantly outperformed free Campto in hypoxia p < 0.05 (50 μg/mL), p < 0.001 (100 μg/mL), and p < 0.001 (250 μg/mL). 250 μM H2
ROS play a complex and somewhat contradictory role in PC. Depending on their concentration within cells, ROS can either support or suppress cancer progression. At low to moderate levels, ROS tend to promote cancer by activating oncogenic signaling pathways that encourage cell growth, survival, and even the ability to spread. However, when ROS levels rise too high, they cause oxidative stress to the cells, which ultimately result in oxidative and mitochondrial damage, leading to cell death through apoptosis.^54^ Next, we evaluated the ability of both free Campto and PD-Campto-CTT1298 to generate ROS in PC3-PIP and C4–2B PC cells (Figure 8C). Using a DCFDA-based confocal fluorescence assay at a concentration of 250 μg/mL, we observed that free Campto induced minimal ROS production, as shown by decreased DCF fluorescence compared to the strong signal from hydrogen peroxide (H2O2), which was used as a positive control. In contrast, PD-Campto-CTT1298 triggered a significant increase in the level of ROS, evident from the intense cytoplasmic DCF fluorescence, nearly matching that of H2O2. These results suggest that PD-Campto-CTT1298 causes substantial oxidative stress in PC cells, which may contribute to cell death through apoptosis or autophagy.
Campto is known to activate autophagy via DNA damage-induced stress signaling pathways, often involving p53 and mTOR suppression.^55^ In PC models, Campto-induced autophagy has been observed to intersect with apoptosis.^56^ To evaluate autophagy in PC3-PIP and C4–2B cells, a CYTO-ID autophagy detection assay was used following treatment with free Campto or PD-Campto-CTT1298. In PC3-PIP cells (Figures 8D and S33), the untreated control exhibited a baseline fluorescence intensity. Treatment with 1 μM rapamycin and 25 μM chloroquine significantly elevated autophagic vesicle accumulation (MFI: ~3809 and ~5714, respectively; p < 0.001), validating the experiment. Free Campto induced a modest increase at 50 μg/mL (MFI: ~1035; p < 0.001 vs untreated), but higher concentrations of 100 μg/mL (MFI: ~409) and 250 μg/mL (MFI: ~390) showed no significant difference from baseline. In contrast, PD-Campto-CTT1298 elicited significantly higher MFI values at 50 and 100 μg/mL (MFI: ~1314 and ~1086; p < 0.001 vs untreated), while 250 μg/mL remained elevated compared to untreated (MFI: ~765; p < 0.01). Similar trends were observed in C4–2B cells (Figures 8E and S34), where rapamycin and chloroquine again produced strong fluorescent signals (MFI: ~4584 and ~9052; p < 0.05 and p < 0.01 vs untreated). Free Campto induced a peak response at 50 μg/mL (MFI: ~1193; p < 0.001 vs untreated), which declined at 100 and 250 μg/mL (MFI: ~777 and ~301; p < 0.01 and ns vs untreated). In contrast, PD-Campto-CTT1298 resulted in significantly higher MFI values at all doses tested (MFI: ~1443, ~1126, and ~847 for 50, 100, and 250 μg/mL; p < 0.001, p < 0.05, and p < 0.01 respectively vs untreated), with consistently greater autophagic signals compared to free Campto. Overall, PD-Campto-CTT1298 consistently induced stronger and more sustained autophagic responses than free Campto across all concentrations of both cell lines.
To investigate the antiangiogenic potential of PD-Campto-CTT1298, VEGF-A secretion levels were quantified using ELISA from conditioned media collected from PC3-PIP and C4–2B cells treated with increasing concentrations of either free Campto or PD-Campto-CTT1298. After 24 h of treatment, VEGF-A absorbance was measured at 450 nm and normalized to untreated cells. In PC3-PIP cells (Figure 9A), free Campto exhibited only minor reductions, with ~0.87 ± 0.07 at 100 μg/mL and ~0.91 ± 0.06 at 250 μg/mL (ns). In contrast, PD-Campto-CTT1298 treatment resulted in a dose-dependent and statistically significant reduction in VEGF-A secretion. At 100 and 250 μg/mL, absorbance decreased to ~0.48 ± 0.05 (p < 0.01) and ~0.28 ± 0.03 (p < 0.001), respectively, compared to untreated controls. A similar trend was observed in C4–2B cells (Figure 9B) where free Campto-treated groups again showed minimal suppression at all of the concentrations tested (ns). However, PD-Campto-CTT1298 significantly suppressed VEGF-A secretion at all the 50 μg/mL (~0.44 ± 0.03; p < 0.01), 100 μg/mL (~0.34 ± 0.03, p < 0.001) and 250 μg/mL (~0.22 ± 0.02, p < 0.01). These findings support prior studies that link topoisomerase I inhibition to transcriptional downregulation of hypoxia-inducible and angiogenic factors such as VEGF-A through HIF-1α destabilization.^57^ More importantly, this reduction in VEGF-A also correlates with increased levels of apoptosis and mitochondrial dysfunction observed in earlier assays. Overall, these data suggest that PD-Campto-CTT1298 employs a multifaceted anticancer effect, targeting both survival and stress-response pathways in PC cells beyond the DNA damage mechanism.
Pathological angiogenesis is a hallmark of PC progression, contributing to tumor growth, metastasis, and therapeutic resistance by promoting endothelial cell proliferation and elevated secretion of VEGF-A and other angiogenic mediators.^58,59^ In order to evaluate the indirect antiangiogenic effects of PD-Campto-CTT1298, we performed a HUVEC tube formation assay using conditioned media collected from PC3-PIP cells treated with Free Campto (250 μg/mL), PD-Campto-CTT1298 (250 μg/mL), or vehicle control. Suramin was included as a positive control due to its well-characterized ability to inhibit angiogenesis by interfering with fibroblast growth factor (FGF) and VEGF signaling.^60^ As shown in Figures 9C and S35, untreated HUVECs formed extensive tubular networks with high total tube length, junction complexity, and mesh formation. Suramin-treated cells resulted in severely disrupted networks, validating the assay’s performance. Conditioned media from free Campto-treated PC3-PIP cells resulted in only minor reductions in angiogenic features (~2–20%), suggesting a limited effect on pro-angiogenic signaling. In contrast, conditioned media from PD-Campto-CTT1298-treated cells induced significant inhibition of angiogenesis. Quantitative analysis using ImageJ ~58% reduction in total tube length (p < 0.001), ~59% reduction in nodes (p < 0.001), ~59% reduction in junctions (p < 0.001), ~66% reduction in segments (p < 0.001), ~77% reduction in master segments (p < 0.001), ~45% reduction in branches (p < 0.001), and 93% reduction in mesh formation (p < 0.001) demonstrating a statistically significant and comprehensive disruption of endothelial tube formation. To validate whether this antiangiogenic effect was consistent, C4–2B cells were also used to perform the same experiment. Similar to PC3-PIP cells, untreated C4–2B conditioned media supported the same angiogenic features including high tube length, branch complexity, and mesh formation. Likewise, Free Campto-treated C4–2B cells induced only mild reductions in tube formation parameters, indicating limited suppression of pro-angiogenic signaling. The conditioned media from PD-Campto-CTT1298-treated C4–2B cells resulted in a drastic inhibition of angiogenesis. ImageJ quantification showed a ~13% reduction in total tube length (p < 0.001), ~62% reduction in nodes (p < 0.001), ~63% reduction in junctions (p < 0.001), ~66% reduction in segments (p < 0.001), ~50% reduction in master segments (p < 0.001), ~49% reduction in branches (p < 0.001), and ~67% reduction in mesh formations (p < 0.001). These findings are consistent with our VEGF-A ELISA results (Figure 9A), where PD-Campto-CTT1298 significantly reduced VEGF-A secretion in PC3-PIP cells (p < 0.001) and C4–2B cells (p < 0.001), while free Campto showed a negligible impact. The enhanced antiangiogenic response of PD-Campto-CTT1298 may be attributed to its improved cellular uptake, sustained release, and induction of apoptosis, as evidenced by caspase-3/7 activation, mitochondrial depolarization, and Annexin V/PI staining reported in previous sections. Together, these results demonstrate that PD-Campto-CTT1298 not only reduces direct angiogenic signaling but also disrupts tumor-secreted pro-angiogenic factors critical to the PC microenvironment.
To evaluate the potential benefit of combining Olaparib (OP) with PD-Campto-CTT1298, PC3-PIP and C4–2B cells were incubated with either OP alone or in a 1 combination with PD-Campto-CTT1298 across a dose range of 50–500 μg/mL. Cell viability was assessed after 48 h using the CellTiter-Glo assay and notably, across all tested concentrations, the combination treatment significantly reduced cell viability compared to either Free OP or PD-Campto-CTT1298 monotherapy (Figure 10 A,B). In PC3-PIP cells, cotreatment resulted in a marked decrease in viability at 50 μg/mL (~54.7%) compared to OP alone (~84.6%, p < 0.01) and PD-Campto-CTT1298 alone (p < 0.01). The cytotoxic effect was correlated to a dose-dependent relationship. At 500 μg/mL, the combination showed the greatest reduction in viability (~31.4%), significantly lower than either Free OP (~65%, p < 0.001) or PD-Campto-CTT1298 (p < 0.01). Similarly, in C4–2B cells, cotreatment consistently outperformed monotherapies. At 500 μg/mL, the combination therapy led to the most significant reduction (~29.7%) relative to Free OP (~65.4%, p < 0.01) and PD-Campto-CTT1298 (p < 0.01). The enhanced cytotoxicity observed with the combination is consistent with previous reports demonstrating the synergistic antitumor effects of Olaparib with Campto derivatives in epithelial cancers.^61^ Co-treatment strategies that combine PARP inhibition with DNA-damaging agents have been shown to suppress genome instability and drug resistance phenotypes more effectively.^62^ This synergy is likely driven by the dual consequence of Campto inducing DNA breaks via Topoisomerase I inhibition, while OP prevents effective DNA repair by blocking PARP activity.^63^ Synergy was quantitatively assessed using three independent Chou-Talalay Combination Index (CI), Bliss Synergy Analysis, and Highest Single Agent (HSA) (Figure 10C). These well-established methodologies help distinguish between additive and synergistic effects of drug combinations in vitro.^64^ The Chou–Talalay method applies median-effect analysis to characterize drug interactions and define CI values for synergy, additivity, or antagonism.^65^ Bliss Synergy Analysis provides a probabilistic framework for evaluating synergy, while the HSA model compares the combination effect to that of the more potent monotherapy.^66,67^ Synergy analysis revealed strong synergistic effects of the combination treatment in PC3-PIP cells, particularly at lower doses. In C4–2B cells, synergy was more modest and dose-dependent with diminishing effects at higher concentrations. Overall, the combination was more effective and synergistic across most of the concentrations tested. Overall, these findings highlight the enhanced therapeutic efficacy of PD-Campto-CTT1298 when combined with Free OP, particularly in PSMA-expressing PC cells. The synergistic reduction in viability supports the rationale for combining targeted drug delivery systems with DNA repair inhibitors to overcome resistance mechanisms and maximize tumor cytotoxicity.
In this study, we successfully synthesized and characterized PD-Campto-CTT1298, a Camptothecin-loaded PAMAM dendrimer conjugated with the PSMA-targeting ligand CTT1298, using CuAAC and SPAAC click chemistry. The conjugate exhibited high aqueous solubility, excellent batch-to-batch reproducibility, and pH- and esterase-responsive drug release. The modular design allowed for the precise and reproducible attachment of both therapeutic and targeting moieties without compromising structural integrity. Compared to free Campto, the dendrimer demonstrated superior formulation stability and improved aqueous solubility at concentrations suitable for in vitro efficacy studies. While in vivo studies are needed to establish pharmacologically and clinically relevant dose ranges, the enhanced therapeutic profile suggests the potential for transitional utility. Using a Cy5-labeled derivative, we demonstrated enhanced and time-dependent uptake in PSMA-expressing PC cells (PC3-PIP and C4–2B). Mechanistic studies confirmed PSMA-mediated uptake via clathrin-dependent endocytosis and subsequent trafficking to lysosomes, as evidenced by blocking experiments and confocal imaging. Functionally, PD-Campto-CTT1298 exhibited potent, dose-dependent cytotoxicity and proliferation inhibition, surpassing the activity of free Campto. We observed hallmark features of mitochondria-mediated apoptosis, including mitochondrial membrane depolarization, caspase-3/7 activation, and ROS generation. Furthermore, PD-Campto-CTT1298 reduced VEGF-A secretion and disrupted endothelial tube formation, indicating antiangiogenic potential. Autophagy was also induced, suggesting autophagic flux as a resistance mechanism that can be therapeutically exploited. Furthermore, PD-Campto-CTT1298 reduced VEGF-A secretion and disrupted endothelial tube formation, indicating antiangiogenic potential. Finally, combination treatment with free Olaparib yielded synergistic effects, reinforcing the therapeutic potential of PD-Campto-CTT1298 as both a standalone and combinatorial nanotherapeutic. The integration of targeted delivery sustained intracellular release, and modulation of multiple tumor survival pathways reflects the platform’s comprehensive therapeutic design. This mechanistic versatility may help overcome key barriers in PC treatment, including resistance and tumor heterogeneity. Together, these findings highlight the value of rationally designed dendrimer-based systems for achieving tumor-specific delivery and multifunctional therapeutic action ultimately for targeted PC therapy.
The Supporting Information is available free of charge at https://pubs.acs.org/doi/10.1021/acsanm.5c03528.
Supplementary figures for NMR Spectral data, HPLC chromatogram, mass spectra, DLS size/zeta potential distribution graphs of the intermediates and dendrimers, flow cytometry data, cytotoxicity analysis, apoptosis assay, caspase activity assay, hypoxia, autophagy, and angiogenesis assay (PDF)