Delivery of Multifunctional Microspheres via Intravesical Instillation for Bladder Carcinoma: Therapeutic Potential and Mechanistic Insights

ABSTRACT

Despite the use of standard intravesical instillation after surgery for non‐muscle invasive bladder cancer (NMIBC), current drugs fail to eradicate residual tumor cells and are associated with high recurrence rates. To address this unmet clinical need, we synthesized tumor‐targeted poly(1‐carboxy‐3,3,3‐trimethylpropyl methacrylate) (PCTM) microspheres for intravesical delivery using microfluidic and click chemistry. Systematic in vitro and in vivo evaluations demonstrated that the PCTM microspheres exhibit excellent bladder mucosal adhesion, deep tissue penetration, and tumor‐specific targeting. Notably, PCTM exerted potent antitumor activity in a variety of bladder cancer cell lines and in orthotopic and ectopic models of urinary bladder cancer, with no serious side effects detected in the peripheral vital organs of mice. Mechanistic studies integrating RNA sequencing, flow cytometry, and western blot analysis showed that PCTM inhibited the proliferation and progression of bladder cancer cells by downregulating CDC42 and HCFC1 and indirectly inhibiting the expression of the cell‐cycle protein A1, which subsequently induced S‐phase arrest of mitosis. Taken together, these findings suggest that the PCTM microspheres may be a promising intravesical therapeutic agent for bladder cancer tumor eradication, thus providing a transformative strategy to significantly improve the prognosis of patients with NMIBC.

PCTM microspheres were successfully synthesized using microfluidic and click chemistry techniques.In Vivo and in vitro experiments have shown that PCTM microspheres have obvious targeting, adhesion, membrane penetration and anti‐tumor properties for bladder tumors.The antitumor mechanism of PCTM microspheres is to indirectly induce S‐stage blockage of cell division by downregulating CDC42 and HCFC1 proteins, which in turn leads to apoptosis of tumor cells.

1. Introduction

Bladder tumors are the most common and lethal cancers in urology [1, 2], 75% of which are non‐muscle invasive [3]. Despite routine postoperative bladder perfusion therapy, the recurrence rate of bladder cancer two years after surgery is as high as 31%–78% [4]. This is attributed to several reasons: the urine in the patient's bladder constantly dilutes the bladder perfusion drugs, preventing them from reaching the effective antitumor concentration [5]; the infused drugs may fail to adhere to the surface of bladder tumors and can be rapidly excreted in urine; most infusion drugs lack specific targeting capabilities for bladder tumors and may cause collateral damage; the protective effect of the bladder–urinary tract epithelial barrier significantly hinders the penetration of the drug into the deeper layers, thereby impeding its efficacy in achieving tumor‐killing effects; and some bladder perfusion drugs are prone to inflammatory reactions in the urinary tract, making the scheduled course of treatment difficult to complete.

Contemporary research has acknowledged these shortcomings and achieved certain improvements. For example, researchers prepared composite microspheres encapsulating poloxamer gel and gemcitabine HCl to enhance drug adhesion to the bladder mucosa [6]. These composite microspheres demonstrate prolonged intravesical retention times and strong anti‐bladder‐tumor effects. Additionally, researchers developed poly(lactic acid) composite microspheres encapsulating hydroxycamptothecin, which demonstrated significant sustained‐release capabilities and antitumor properties during bladder instillation therapy (BIT) [7]. This approach addresses the issue of insufficient effective drug concentrations during treatment. Yin et al. synthesized magnetic hydrogel microspheres encapsulating doxorubicin [8]. Through the synergistic effects of the controlled magnetic fields and regulated urinary pH, sustained effective drug concentrations were achieved during bladder instillation. However, these studies did not comprehensively address the challenges associated with BIT. Currently, there is an urgent need to develop a bladder perfusion drug that not only targets bladder tumors and achieves adhesion, penetration, and antitumor functions, but is also free of significant systemic toxic side effects.

In view of this, we synthesized multifunctional poly(1‐carboxy‐3,3,3‐trimethylpropyl methacrylate) (PCTM) microspheres containing polylactic acid (PLA), Cy5, tannic acid (TA), and polypeptides R11 and M3‐2K using microfluidic and click chemistry techniques and applied them to bladder perfusion therapy to kill tumor cells and reduce recurrence. PLA is a safe and nontoxic degradable protein suitable for carrier delivery and locally controlled drug release [9]. TA has bioadhesive properties that promote wound healing [10]. We applied TA to the bladder mucosa for the first time to increase the adhesion of PCTM microspheres. To observe the localization of and changes in the PCTM microspheres during the experiment, we introduced Cy5 cell membrane markers [11]. Membrane‐penetrating peptides are peptide sequences that can rapidly cross cell membranes and deliver a wide range of substances within the cells, including proteins and nucleic acids. R11 is an 11‐peptide arginine membrane‐penetrating peptide that not only penetrates the cell membrane but also targets bladder tumors [12]. In recent years, the peptide amino acid M3‐2K [13], which exhibits extremely high antitumor activity, was discovered using phage technology to have extremely low toxicity, but it has not yet been applied to solid tumors. We incorporated M3‐2K and R11 polypeptides into a novel complex protein using click chemistry for the first time and analyzed its tumor‐targeting and antitumor abilities in vivo and in vitro.

Bladder perfusion is the standard treatment modality for non‐muscle‐invasive bladder cancer (NMIBC). In this study, the PCTM microsphere–enhanced adhesion and permeability to the bladder mucosa, as well as the microsphere‐targeting and antitumor effects on bladder tumor cells, were verified through a series of cellular experiments and a nude mouse model of bladder cancer. Its biocompatibility was verified by examining pathological sections of other important organs in the whole body and hematological indicators. Further studies revealed that PCTM microspheres exert their antitumor effects by downregulating the expression of several cell‐cycle proteins (e.g., CDC42, HCFC1, cyclin A1), which in turn blocks the S‐phase of the tumor cell cycle and leads to apoptotic cell death [14, 15]. These results demonstrate that bladder perfusion is highly promising for eradicating bladder tumor cells, reducing the recurrence rate and improving prognosis (Scheme 1).

SCHEME 1.

Synthesis process and mechanism of action of poly(1‐carboxy‐3,3,3‐trimethylpropyl methacrylate) (PCTM) microspheres. (A) Synthesis and characteristic functions of PCTM microspheres. (B) Mechanism of action of PCTM microspheres against bladder tumor. (C) Schematic showing bladder perfusion of the PCTM microspheres.

2. Results and Discussion

2.1. Preparation and Characterization of PCTM Microspheres

For the intravesical instillation of drugs to radically treat bladder cancer, we designed novel microspheres (Figure 1A) with the intended characteristics of targeting, membrane penetration, antitumor activity, and biocompatibility. PLA gradually degrades into nontoxic CO₂ and H₂O through hydrolysis or enzymatic degradation at degradation rates generally ranging from a few days to a few months, which meets the requirements for the sustained release of bladder perfusion drugs. In addition, microfluidic technology can be used to accurately control the size and appearance of nanoparticles and improve the controllability of drug loading and release. Therefore, this technology was used to successfully prepare PLA‐Cy5 microspheres [16]. TA mainly binds to PLA through hydrogen bonding; its hydrophobicity stabilizes the microspheres and may give rise to adhesiveness or anti‐inflammatory effects [17]. The targeting and membrane‐permeability mechanisms of R11 potentially involve two factors: one on hand, R11 has multiple arginine residues, and these positively charged residues enable it to interact with the negatively charged components of the cell membrane; on the other hand, the ITGB1 locus is highly expressed in bladder cancer cells, and the R11 peptide achieves its targeting function by preferentially binding to the ITGB1 gene [18]. Previous studies have shown that M3‐2K can target the MDM2 and MDMX proteins, leading to the activation of the oncoprotein p53, which produces anti‐malignant glioma effects [13]. Considering that oncoprotein P53 closely influences the prognosis of bladder cancer, we hypothesized that M3‐2K may exert anti‐bladder‐cancer effects. Therefore, we chemically synthesized a R11‐M3‐2K/fluorescein isothiocyanate (FITC) complex using click chemistry. FITC is a fluorescent dye that exhibits bright green fluorescence under a microscope and can be applied to cell membranes for intracellular tracing.

FIGURE 1.

Preparation process and general characterization of PCTM. (A) Step‐by‐step synthesis of PCTM microspheres. (B) Diameter and surface roughness of the four groups of microspheres, observed using SEM. (C,D) Surface roughness of the four groups of microspheres analyzed by atomic force microscopy (AFM). (F) high‐performance liquid chromatography (HPLC) analysis of RM peptide. (G) X‐ray photoelectron spectroscopy (XPS) spectra of the PTCM microspheres. (H) Infrared (IR) spectra showing the main functional groups in the four groups of microspheres.

The electron microscopy images revealed that the average particle size of the four groups of microspheres gradually increased from 56.23 ± 0.61 to 235.60 ± 41.48 µm. The four groups of microspheres were spherical, and the surface was smooth, as expected, based on the microfluidic technology preparation. With the addition of TA and R11‐M3‐2K (RM) peptides, the surface roughness of the microspheres gradually increased (Figure 1B). Atomic force microscopy (AFM) was used to confirm that the surface roughness of the four groups of microspheres gradually increased, and the PCTM group exhibited the highest roughness (Figure 1C); quantitative analysis yielded the same result (Figure 1D). In addition, we examined the surface charges of the four groups of microspheres. The surface of the P group was negatively charged, possibly originating from the ester group (–COO–) of PLA or the carboxylic acid group (–COOH) produced by microdegradation, whereas the PC, PCT, and PCTM groups showed an obvious positive charge, possibly owing to the polyphenolic hydroxyl group of TA and modification of the targeted peptide guanidinium group (Figure 1E). The positive charge on the surface of the PCTM microspheres can target negatively charged bladder cancer cells via electrostatic adsorption while promoting receptor‐mediated endocytosis. The RM group exhibited one major peak, indicating that the target peptide was successfully coupled (Figure 1F). The four groups of microspheres were further analyzed using X‐ray photoelectron spectroscopy (XPS; Figure 1G). In the C 1s spectrum of the P group, the main peaks corresponded to C–C (284.8 eV) and C–O (286.9 eV) bonds, consistent with the PLA structure; for the PC/PCT/PCTM group, a C = C peak (284.4 eV, the aromatic ring from Cy5) and an additional O–CO peak (289.1 eV, ester bond of tannins) emerged. In the N 1s spectrum of the PCTM group, peaks corresponding to Ph–NH₂ (399.6 eV) and C = NH (400.9 eV) were apparent, confirming the successful modification of RM. In the O 1s spectrum of the PCT/PCTM group, an ArOH peak (533.5 eV) also appeared, indicating successful incorporation of the polyphenol hydroxyl group of TA and confirming that the four microsphere‐specific groups were successfully loaded into the composite microspheres. The infrared (IR) spectra of the four groups of microspheres are shown in Figure 1H. The strong peak at 1750 cm⁻¹ in the spectrum of the P group corresponds to the C = O ester bond of PLA, whereas the weak peak at 1650 cm⁻¹ in the spectrum of the PC group originates from the C = C aryl ring of Cy5. The 3300–3500 cm⁻¹ peaks in the PCT spectrum indicate that tannins were coordinated to the surface of PLA via hydrogen bonding. In the spectrum of the PCTM group, the intensity of the 1650 cm⁻¹ peak was high, possibly owing to the superposition of the amide bond (N–H) bending vibration of RM, confirming covalent coupling of the peptide segment. The spectra not only confirm the successful preparation of the composite microsphere PCTM but also show that the key structures were maintained.

2.2. Characterization of RM Peptide

The RM peptides were synthesized and characterized. First, the molecular formulae of the A, D, and B chains (Figures S1–S3) and that of the RM peptide (ADB chain) (Figure 2A) were determined using ChemDraw software. Next, we simulated moment diagrams of the structures of the ADB, A, D, and B chains at 0, 20, and 100 ns. Figure 2B and Figure S4 (Supporting Information) show that the four peptide chains start to adjust their structure at 0 ns and reach a relatively stable state at 100 ns, which lays the foundation for their comprehensive function.

FIGURE 2.

Structural characterization of RM. (A) Chemical structure of RM. (B) Structural state of ADB, A, D, and B chains at 0, 20, and 100 ns. (C) Root mean square deviation (RMSD) of ADB, A, D, and B chains over time. (D–G) Solvent accessibility surface area (SASA) of ADB, A, D, and B chains over time. (H) Abundant weak interactions within the ADB chain: hydrogen bonds between (i) the S atom and the dopa hydroxyl group, (ii) the C‐terminal carboxyl group and arginine, (iii) glutamine and the amide bond on the backbone, and (iv) glutamic acid and neighboring glutamine and arginine. (I) Isosurface plot of the electrostatic potential distribution of ADB chains on the van der Waals surface of the molecule. (J) Regional distribution of different charges of the ADB chains. (K,L) Specific atomic distances and hydrogen bonding in ADB chain over time. (M) 3D diagram showing the hydrogen bonds between ADB chains.

We then introduced the root mean square deviation (RMSD) index, which indicates the deviation of the conformation at a certain time point from the target conformation of all atoms, thus serving as an important basis for measuring the stability of the system. The RMSD curves of the four groups of peptide chains showed that the ADB and A chains reached relatively stable states after 30 ns. In order of stability, the D chain was the strongest, followed by the B chain, while the stabilities of the A and ADB chains were similar (Figure 2C). The solvent accessibility surface area (SASA) is an important index for describing the hydrophobicity of proteins; a low SASA value indicates that the contact area between the protein and water is small, the hydrophobic core formed by protein folding is more compact, and the structure is more stable. Analysis of the four groups of peptide chains (Figure 2D‐G) showed that the SASA of the A, B, and ADB chains gradually decreased and stabilized over time, indicating that the overall hydrophobicity increased, and the structure was more stable. The hydrogen bonds and relative distances between atoms within the molecular chains of the A, D, and B chains were analyzed separately (Figures S5–S10, Supporting Information). The number of hydrogen bonds and distances between the nitrogen atoms in the ARG residues and the amino nitrogen atoms on the LYS of the A chain stabilized over time. The D chain formed fewer hydrogen bonds, the B chain increased the number of hydrogen bonds owing to folding, and the distances between the DOPA carbons and FITC carbons were relatively stable. A further analysis of the secondary structures of the A chain (Figure S11, Supporting Information) and ADB chain (Figure S12, Supporting Information) revealed that the main region of the A chain maintains a stable state of switching back and forth between the turns and 3‐helices and the main region of the ADB‐chain ultimately maintains the α‐helix secondary structure, which are verified in the simulated moment diagrams.

To better understand the spatial conformation and charge distribution of the ADB chain, density functional theory was used to demonstrate the abundant weak interactions within the molecule, which are the basis for the formation of stable conformations (Figure 2H). In summary, the stabilizing structure of the ADB chain relies mainly on hydrogen bonding, supplemented by van der Waals bonding. Isosurface maps of the distribution of ADB‐chain electrostatic sites on the van der Waals surface of the molecule (Figure 2I) and a ball‐and‐stick model of the molecular structure of the ADB chain (Figure 2J, Figure S13, Supporting Information) reveal that the noncovalent bonds formed a specific spatial configuration to maintain structural stability. Next, the stability of the ADB chain was corroborated by the relative stabilization of the interatomic distances between the carbon atoms on the benzene ring at the end of FITC and the amino nitrogen atoms on LYS after fine‐tuning (Figure 2K). The number of hydrogen bonds formed by ADB gradually stabilized in the range of 20–22, indicating that the structure of the ADB polypeptide was fine‐tuned during folding, resulting in subtle changes in the number of hydrogen bonds (Figure 2L). The stability of the ADB structure was demonstrated using the Artoon model. The docking score of ADB to ADB was –304.11, and the confidence score was 0.9562, which proved that the complex model obtained by docking had a high confidence level (Figure S14, Supporting Information). The three‐dimensional (3D) diagram of hydrogen bonding between ADBs (Figure 2M) was analyzed using PyMOL software, and a two‐dimensional (2D diagram) of ADB interactions (Figure S15, Supporting Information) was constructed using Ligplot +v2.2.8, both of which showed that the ADB polypeptides formed stable molecular structures owing to noncovalent bonding interactions. Furthermore, high‐performance liquid chromatography (HPLC) and mass spectrometry (MS) analyses confirmed that RM peptide synthesis was successful and achieved a purity of 95.90% (Figure S16, Supporting Information). Finally, the loading efficiency and release profiles of RM peptides from the PCTM microspheres were determined. Figure S17 (Supporting Information) shows that the PCTM microspheres exhibited a loading efficiency of up to 20.76% for the RM peptide. This is likely attributable to the vast internal space and complex porous structure within the micron‐sized microspheres, which can accommodate a large number of drug molecules (e.g., TA and peptide M3‐2K). Figure S18 (Supporting Information) shows that the RM peptide underwent a burst release effect during the initial phase (day 1), followed by a sustained release phase (days 1–14), during which the cumulative release reached 77.50%. Finally, the slow‐release phase (days 14–20) resulted in a cumulative release rate of 81.77%. Thus, the PCTM microspheres are ideal drug‐delivery platforms for bladder instillation.

2.3. Targeting and Membrane Penetration Verification of RM Peptide in Bladder Tumor Cells

Next, we conducted a series of in vivo and in vitro experiments to fully demonstrate the combined advantages of the PCTM microspheres, including their adhesion and permeability to the bladder mucosa, targeting ability, antitumor effects on bladder tumors, and biocompatibility (Figure 3A). Because the substances exerting membrane‐penetrating, targeting functions in PCTM microspheres are likely to be gradually released RM peptides, we used 5 µg/mL PCTM microspheres equivalently loaded with 1 µg/mL free RM peptide for direct experimental observation, which avoids interference with the complex release kinetics of the delivery system. First, the fluorescence intensities were measured 3, 6, and 12 h after T24, T5637, and SV‐HUC‐1 cells were treated with 1 µg mL⁻¹ RM peptide (Figure 3B‐D). We found that the fluorescence intensity of the SV‐HUC‐1 cells was weaker 3 and 6 h after treatment. Compared with the SV‐HUC‐1 cells, the T24 and T5637 cells had significantly higher fluorescence intensities at 3, 6, and 12 h (Figure 3E). Compared with the fluorescence intensities of the T24 and T5637 cells 3 h after treatment, treatment at 6 and 12 h showed a slow increase in fluorescence intensity (Figure 3F,G). These results reveal that the RM peptide has no obvious function in targeting SV‐HUC‐1 cells, but has an obvious and continuous function in targeting T24 and T5637 cells, which creates favorable conditions for antitumor activity.

FIGURE 3.

Membrane penetration and targeting functions of RM peptides. (A) Molecular formulae and corresponding functions of R11, M3‐2k, and tannic acid. (B–G) RM‐targeting experiments in T24, T5637, and SV‐HUC‐1 cells and application of ImageJ software for quantitative fluorescence analysis. (H–N) RM peptides in T24, T5637, and SV‐HUC‐1 cells for membrane‐penetration assay and application of ImageJ for quantitative fluorescence analysis.

Subsequently, the membrane‐penetration status of the RM peptide in SV‐HUC‐1, T24, and T5637 cells was evaluated (Figure 3H–J). DiI‐labeled cell membranes showed red fluorescence, FITC‐labeled RM peptides showed green fluorescence, and the overlay was a superimposed image of the two types of fluorescence. At 3 h, green fluorescence was already evident in the T5637 and T24 cells, indicating that the RM peptide started to penetrate the cell membrane into the cytoplasm. As the coculture time was extended to 6, 9, and 12 h, the distribution of the RM peptide became more widespread and the fluorescence intensity gradually increased in the T5637 and T24 cells, indicating that the RM peptide was gradually taken up by the cells and might be transported by or accumulated in the cells. By contrast, the green fluorescence intensity of the SVHUC‐1 cells was significantly weaker and gradually decreased over time. Further quantitative analysis showed that the FITC fluorescence intensities of the T5637 and T24 cells were significantly stronger than those of the SVHUC‐1 cells at 3, 6, 9, and 12 h. The FITC fluorescence intensities of the T5637 and T24 cells tended to increase over time, whereas that of the SVHUC‐1 cells was persistently weak (Figure 3K). This demonstrated that the membrane‐penetration efficiency of the RM peptide in the T5637 and T24 cells was much higher than that in the SVHUC‐1 cells. Figure 3L shows the changes in the FITC fluorescence intensity of the same cells at different time points. The fluorescence intensity of SVHUC‐1 cells was consistently weaker at 3, 6, and 9 h, with an even weaker state after 9 h. By contrast, the 9‐ and 12‐h fluorescence intensities of the T5637 and T24 cells were much higher than those at 3 h. This indicates that the RM peptides are more likely to pass through the membranes of tumor cells and have a lower membrane‐penetrating effect on normal cells. Therefore, the RM peptide selectively acts on tumor cells and reduces damage to normal tissues.

Determining the trend in FITC fluorescence intensity of the three cell types during the experimental process elucidated the dynamic membrane‐penetration process of the RM peptide into the cells. As shown in Figure 3M, the FITC fluorescence intensities of the T5637 and T24 cells gradually increased after 3 h and reached a high level at ≈12 h, whereas the fluorescence intensity of the SVHUC‐1 cells did not change significantly and remained at a low level throughout the time scale. These results indicate that the increased fluorescence intensity of the RM peptides during membrane penetration into tumor cells is mainly attributed to the progressive release of RM peptides from the PCTM microspheres over time. The trends in the DiI fluorescence intensity of the three cell types in the same time range are shown in Figure 3N; the fluorescence intensity changes in the DiI‐labeled cell membranes indirectly reflect the integrity and stability of cell membranes. As shown in the figure, the DiI fluorescence intensity of the three cell types changed relatively little during the experimental period, and was similar at each time point. This indicates that the integrity and stability of the cell membrane were not significantly affected by the membrane penetration of the RM peptide. A possible explanation is that the membrane‐penetrating process of the RM peptide might proceed through a normal cellular uptake mechanism rather than entering the cells by destruction of the cell membrane.

2.4. Antitumor Properties of PCTM Microspheres in Bladder Tumor Cells

Live/dead staining experiments were performed to evaluate the antitumor properties of the PCTM microspheres. Fluorescence microscopy images of the SVHUC‐1, T5637, and T24 cells treated with the P, PC, PCT, or PCTM microspheres are shown in Figure 4A–C. The low‐intensity red fluorescence in the SVHUC‐1 cells treated with the P, PC, PCT, and PCTM microspheres indicate that the apoptosis‐inducing effect of these four groups of microspheres on normal cells was weak. In the T5637 and T24 cells, the intensity of red fluorescence gradually increased, and the proportion of apoptotic cells increased significantly, indicating that the PCTM microspheres had a strong apoptosis‐inducing effect on tumor cells. Subsequently, the results of four subsequent sets of microspheres treating three types of cells are shown: the cell‐death rates of SVHUC‐1 cells treated with the four groups of microspheres were all considerably low, with no obvious difference, whereas the death rates of the T5637 and T24 cells in the PCTM group clearly increased (Figure 4D), confirming that the PCTM microspheres were cytotoxic to tumor cells and induced apoptosis. Subsequently, the viabilities of the SVHUC‐1, T5637, and T24 cells after treatment with varying concentrations of the PCTM microspheres were determined using a ck‐8 kit (Figure 4E). With the gradual increase in PCTM microsphere concentration, the viability of the SVHUC‐1 cells was less affected, while those of the T5637 and T24 cells decreased significantly, particularly at ≥5 µg mL⁻¹. These experimental results indicate that the PCTM microspheres are selectively cytotoxic to tumor cells and induce tumor cell death, providing an important experimental basis for the targeted therapy of bladder cancer. The dose‐dependent inhibitory effect of the PCTM microspheres on tumor cells also indicate that the therapeutic effect could be optimized by adjusting the drug concentration in clinical applications to reduce side effects on normal tissues.

FIGURE 4.

Evaluation of antitumor properties of poly(1‐carboxy‐3,3,3‐trimethylpropyl methacrylate) (PCTM) microspheres against bladder cancer cells. (A–D) Four groups of microspheres (P, PC, PCT, and PCTM) applied to live–dead experiments and SVHUC‐1, T5637, and T24 cells. (E) CCK‐8 experiments with PCTM microspheres in SVHUC‐1, T5637, and T24 cells. (F–K) Four groups of microspheres (P, PC, PCT and PCTM) used for cell‐scratching experiments and analysis of SVHUC‐1, T5637, and T24 cells.

Next, we investigated whether the PCTM microspheres inhibited tumor cell migration in a cell scratch experiment. A concentration of 5 µg mL⁻¹ of each type of microsphere (P, PC, PCT, and PCTM) was used to treat T5637 and T24 cells separately, and the changes in the widths of the scratches were determined at 0, 6, 12, and 24 h. The results showed that the scratches on T5637 cells in the P, PC, and PCT groups gradually healed over time, whereas those in the PCTM group still had a large, unhealed area at 24 h (Figure 4F). Quantitative analysis showed that the migration areas of the P, PC, and PCT groups were significantly increased at 6, 12, and 24 h relative to those of the PCTM group (Figure 4G), and the migration distances of the P, PC, and PCT groups were significantly increased at 12 and 24 h relative to those of the PCTM group (Figure 4H). Similarly, images of the microsphere‐treated T24 cells at 0, 6, 12, and 24 h showed that over time, the scratched areas in all groups except the PCTM group were filled with a large number of cells in a shorter period (Figure 4I). Quantitative analysis showed that the migration area and distance of the P, PC, and PCT groups at 6, 12, and 24 h were significantly higher than those of the PCTM group (Figure 4J,K).

The antitumor properties of the PCTM microspheres were validated using a cell clone formation assay (Figure S19, Supporting Information). SVHUC‐1, T5637, and T24 cells were each cultured in media separately containing 5 µg mL⁻¹ P, PC, PCT, and PCTM, and the number of cell colonies were counted after 10 days. The T5637 and T24 cells showed strong colony‐forming ability and high cell numbers in the P, PC, and PCT groups, whereas the colony‐forming ability was significantly weaker in the PCTM group, and the cell number was also lower. These results confirmed that the PCTM microspheres not only exerted concentration‐dependent cytotoxic effects on tumor cells but also inhibited tumor metastasis. They also exhibit minimal toxic side effects in normal cells. These findings strongly support the safety and efficacy of the PCTM microspheres for the treatment of bladder tumors.

2.5. Potential Bladder Tumor Eradication Mechanism of PCTM Microspheres

To further unravel the potential mechanism of the antitumor activity of PCTM microspheres, we treated T5637 and T24 cells with 5 µg mL⁻¹ PCTM and blank medium, respectively, and then sent them for transcriptome sequencing (Table S1, Supporting Information). After analyzing the sequencing data of the experimental and control groups, volcano plots of the differential genes in the T5637 (Figure 5A) and T24 groups (Figure 5D) were constructed, where the red dots indicate significantly upregulated genes, the blue dots indicate significantly downregulated genes, and the gray dots indicate genes with no statistically significant differences. In both plots, the expression of HCFC1 and CDC42, both of which play important roles in cell cycle regulation, was significantly downregulated. In the heatmaps of the top 20 genes upregulated and downregulated by sequencing in the two groups (Figure 5B,E), a large number of genes involved in cell‐cycle regulation (marked in red) were identified. Further pathway analysis showed that these differential genes were mainly concentrated in the cell cycle, apoptosis, and autophagy pathways, with cell cycle ranking first (Figure 5C,F). This indicates that PTCM microspheres exert their antitumor effects primarily through cell‐cycle regulation. Flow cytometry, apoptosis, and cell‐cycle assays were performed. First, the apoptosis results show that early apoptosis (Q3) and late apoptosis (Q2) of SVHUC‐1 cells increased slowly over time, and the total apoptosis rate of Q was 23.51% after 12 h. By contrast, early apoptosis (Q3) and late apoptosis (Q2) increased rapidly in the T5637 and T24 cell lines, with total apoptosis rates of 77.9% and 82.5%, respectively, after 12 h (Figure 5G). This indicates that the PTCM microspheres had specific apoptosis‐inducing effects on bladder tumor cells: the longer the duration of action, the higher the proportion of apoptotic cells, whereas the apoptosis‐inducing effect on normal cell lines was relatively weak. Next, the cell‐cycle results showed that the proportion of the S phase in T5637 cells gradually increased, the proportion of the G0/G1 phase in the T24 cell line gradually decreased, and the proportion of the S phase gradually increased, indicating that the PTCM microspheres may exert their antitumor effects through the G1/S phase (Figure 5I).

FIGURE 5.

Antitumor mechanism of poly(1‐carboxy‐3,3,3‐trimethylpropyl methacrylate) (PCTM) microspheres. (A–C) Differential gene, heat‐map, and pathway analysis of transcriptome sequencing before and after PCTM microsphere treatment of T5637 cells. (D–F) Differential gene, heat‐map, and pathway analysis of transcriptome sequencing before and after PCTM microsphere treatment of T24 cells. (G) Flow cytometric apoptosis detection of PCTM microsphere–treated SVHUC‐1, T5637, and T24 cells. (H) Western blots of critical cell‐cycle proteins. (I) Flow cytometric cycle detection of PCTM microsphere–treated SVHUC‐1, T5637, and T24 cells. (J) Diagram illustrating the antitumor mechanism of PCTM microspheres.

To determine the mechanism by which PTCM microspheres regulate the G1/S phase of the cell cycle, the important proteins reported so far to regulate the G1/S phase were analyzed by western blotting: as the concentration of PCTM microspheres was increased from 0 to 8 µg mL⁻¹, the expression of HCFC1, CDC42, and cyclin A1 gradually declined, whereas internal reference protein (ACTIN) expression remained unchanged (Figure 5H). These results indicate that the mechanism of action of the PTCM microspheres on bladder tumors mainly involved the downregulation of the expression of HCFC1, CDC42, and cyclin A1 proteins, which in turn blocked the G1/S phase of the cell cycle and achieved antitumor effects. Numerous studies have shown that these key proteins regulate cell‐cycle processes. Xiang et al. reported that HCFC1 binds to the –881 to –575 region upstream of the CDC42 transcriptional start site and facilitates CDC42 expression, whereas CDC42 expression decreases when HCFC1 is silenced. Overexpression of CDC42 can rescue the decrease in cyclin A expression caused by HCFC1 silencing, whereas downregulation of CDC42 can lead to a decrease in cyclin A expression through the phosphorylation of E2F by the downstream effector protein PAK1 [14]. Tyagi et al. reported that HCFC1 recruits MLL and SET1 histone H3K4 methyltransferase to the E2F‐responsive promoter, thereby inducing transcriptional activation and indirectly affecting cyclin A1 expression [19]. Ji et al. reported that the cyclin A1–CDK2 complex phosphorylates Rb‐family proteins, deregulates their inhibition of E2F, and activates the S phase of the cell cycle. When cyclin A1 was inhibited, progression from the G1 phase to the S phase of the cell cycle was blocked [20]. In summary, we believe that HCFC1 and CDC42 can act indirectly on cyclin A1, inhibiting DNA replication and regulating the progression of the G1/S phase of the cell cycle, ultimately resulting in tumor‐cell necrosis and apoptosis (Figure 5J).

2.6. Validation of Adhesion and Permeation Function of PCTM Microspheres in Mouse Model of Bladder Cancer

Next, we evaluated whether the PCTM microspheres possessed the adhesion and permeation functions required for bladder perfusion in vivo. First, we treated blank cells, T24 cells, and nude mice with the four microspheres and used near‐infrared (NIR) thermography to determine whether the microspheres could effectively enter the corresponding tissues. As shown in Figure 6A, under NIR thermography, the red microspheres in the PC, PCT, and PCTM groups (i.e., all groups except for the P group) were visible because these groups contained the Cy5 red‐fluorescent dye. NIR thermography showed that the microspheres in all three groups reached the bladder adequately, reliably guaranteeing subsequent bladder perfusion. These three groups of microspheres were perfused into the bladders of the nude mice, and the fluorescence intensity of the bladder samples was observed at 0.5, 2, 6, 12, and 24 h to evaluate the adhesion of the microspheres in each group. As shown in Figure 6B, the fluorescence intensity of the PC group gradually decreased over time, whereas those of the PCT and PCTM groups remained strong. Further semiquantitative analysis using Image J software showed that the fluorescence intensity of the PC group gradually decreased with extended observation time, while the fluorescence intensity of the PCT and PCTM groups peaked at 6 h and remained high (Figure 6D); the fluorescence intensities of the PCT and PCTM groups were significantly higher than those of the PC group at 2, 6, 12, and 24 h (Figure 6E). These results confirm, for the first time, that the adhesion of PCT and PCTM groups to bladder mucosa is significantly enhanced by the presence of tannins. The results of the bladder permeability experiments using the PCTM microspheres showed that the whole bladder permeability of the PCT and PCTM groups continued to increase with observation time, whereas that of the PC group was significantly weaker (Figure 6C). Further quantitative analysis revealed that the PCTM group had the best bladder‐wall permeability of the three microsphere bladder‐perfusion groups (Figure 6F). These results confirm that the PCTM microspheres not only continuously enhanced bladder adhesion but also had excellent bladder‐wall penetrability, thus laying the foundation for the radical treatment of deep bladder tumors in the PCTM group.

FIGURE 6.

Adhesion and permeation function of poly(1‐carboxy‐3,3,3‐trimethylpropyl methacrylate) (PCTM) microspheres in the bladder. (A) Near‐infrared (NIR) thermal imaging of four groups of microspheres in T24 cells and nude mice. (B,D,E) Adhesion experiments and quantitative fluorescence analysis of three groups of microspheres (PC, PCT, and PCTM) in bladder mucosa. (C,F) Permeation experiments and quantitative fluorescence analysis of PCTM microspheres in bladder wall.

2.7. Therapeutic Effect of PCTM Microspheres in Bladder‐Cancer Mice

We constructed an in situ bladder‐cancer nude mouse model to study the antitumor effects of the PCTM microspheres in vivo. In nude mice, cystography can be used to visualize the filling defect of the bladder, which is key evidence for the successful construction of an animal model of bladder cancer (Figure 7A, Figure S20, Supporting Information). Two randomly selected nude mice were incised in the bladder, which allowed the detection of a solid mass in the bladder lumen (Figure 7B). Next, bladder tumor tissue sections were stained with hematoxylin and eosin (H&E) to confirm that the bladder tumor cells had cancer nests and nuclear division (Figure 7C). Immunohistochemistry for caspase‐3 and Ki67 proteins showed that the bladder cancer cells exhibited rapid proliferation and invasion (Figure 7D,E). In summary, we successfully established an in situ bladder‐cancer model using nude mice to lay the foundation for in vivo experiments. Next, 2 mL of phosphate‐buffered saline (PBS) or one of the four groups of microspheres (P, PC, PCT, or PCTM) was injected into the bladder once a week, and the bladder was removed after 4 weeks. We discovered that, except for bladder tumors in the PCTM group, which were significantly reduced, those in the other four groups were significantly enlarged after 4 weeks. These results confirm that the PCTM microspheres exhibited tumor‐inhibitory and tumor‐killing effects in vivo (Figure 7F). Subsequent H&E staining of the bladder tumors in each group (Figure 7G) showed a large number of nuclear isoforms (black arrows), nuclear schizophrenia (green arrows), and extensive cell necrosis (yellow arrows) in the pathological sections of the PBS, P, PC, and PCT groups; however, the PCTM group contained much fewer necrotic cells.

FIGURE 7.

Antitumor effects of poly(1‐carboxy‐3,3,3‐trimethylpropyl methacrylate) (PCTM) microspheres in nude mice with in situ bladder cancer. (A) Cystography after modeling of bladder cancer. (B) Visualization of tumors in the bladder. (C–E) Hematoxylin and eosin (H&E) staining, and immunohistochemical analysis of caspase‐3 and Ki67 in bladder tumors. (F) Bladder‐tumor specimens after 4 weeks of drug treatment in five groups. (G) H&E staining of the bladder tumors in five groups, nuclear isoforms (black arrows), nuclear schizophrenia (green arrows), and cell necrosis (yellow arrows). (H) H&E staining of heart, liver, spleen, lung and kidney after drug treatment in five groups. (I,J) Variation in body weight and tumor volume after 4 weeks of drug treatment in five groups. (K–Q) Alanine aminotransferase (ALT), alanine aminotransferase (AST), urea nitrogen (BUN), creatine kinase (Cr), and creatine kinase (CK) assays in bladder‐cancer nude mice after 4 weeks of treatment with the five groups of drugs.

After treating the five groups of nude mice for 4 weeks, H&E staining of the heart, liver, spleen, lung, kidney, and other organs of bladder‐cancer mice showed that the original tissue cells were clear—none were infiltrated by tumor cells (Figure 7H, Figure S21, Supporting Information). Therefore, the PCTM microspheres demonstrated not only local antitumor effects in vivo but also reliable biosafety. Subsequent analysis showed that the body weights of the five groups of nude mice remained relatively stable during the 4‐week treatment period (Figure 7I). Relative to the volumes of the bladder tumors after 1 week of treatment, the tumor volumes of the PBS, P, PC, and PCT groups increased significantly after 4 weeks of treatment, whereas the tumor volume of the PCTM group was significantly reduced (Figure 7J). This confirms that the PCTM microspheres exhibited significant anti‐bladder‐tumor properties in vivo. Analysis of the blood biochemical indices of the five groups of nude mice showed that the blood indices of alanine aminotransferase, alanine aminotransferase, urea nitrogen, creatine, and creatine kinase in nude mice were within the normal range after 4 weeks of treatment (Figure 7K–Q), confirming the biosafety of the PCTM microspheres.

To more accurately analyze the tumor‐inhibiting process of the PCTM microspheres in vivo, we designed five groups of subcutaneous tumor‐bearing nude mice (Figure S22, Supporting Information); H&E staining confirmed the modeling success (Figure S23, Supporting Information). The five groups of microspheres were administered for 4 weeks when the tumor volume was ≈100 mm (Figure S24, Supporting Information). H&E staining of the heart, liver, spleen, lungs, and kidneys of the nude mice after 4 weeks of treatment showed that the structure of each vital organ was intact, and no tumor or inflammatory cell infiltration was observed (Figure S25, Supporting Information). Except for the PCTM group, in which the subcutaneous tumors decreased significantly after 4 weeks of treatment, the subcutaneous tumor volume gradually increased in the other four groups, and one nude mouse died in each of the PBS and P groups (Figure S26, Supporting Information). The subcutaneous tumor volume in the PCTM group initially increased slightly and then gradually decreased during the treatment period (Figure S27, Supporting Information). The body weights of the five groups of nude mice did not significantly fluctuate during the 4‐week treatment (Figure S28, Supporting Information), implying that no serious side effects led to the loss of body weight in the nude mice. The results of these experiments reaffirm that the PCTM microspheres not only gradually inhibit and kill tumors but also have reliable biosafety.

The size of the PCTM microspheres offers multiple advantages during bladder instillation. Their average diameter of 235.6 µm places them in the large‐sized micrometer‐scale carrier category. These large dimensions imply a vast internal space and a complex porous structure within the microspheres. This allows for the loading of large quantities of drug molecules (e.g., TA, RM peptide). The pore size can be regulated using oil/water‐phase preparation processes to achieve efficient drug loading and controlled drug release. Furthermore, “anchored” adhesion rather than intracellular retention occurs: at 235.6 µm, these microspheres cannot enter individual cells via endocytosis. The microspheres’ vast surface area and surface‐modified adhesive components (e.g., TA, targeting peptide R11) enable them to adhere firmly to tumor tissue surfaces or interstices, analogously to Velcro. This potent adhesion facilitates local retention and prevents rapid urine clearance. The formation of a drug reservoir at the target site enables sustained drug release. In addition, owing to their large size, these microspheres degraded relatively slowly over several months. This represents a significant advantage for enabling sustained and stable drug release. Consequently, effective drug concentrations are maintained locally within the tumor over extended periods, reducing the frequency of bladder instillations and improving treatment compliance.

The current bladder instillation systems directly administer chemotherapeutic drugs (such as gemcitabine or doxorubicin) or immunosuppressants (such as BCG) to the bladder cavity to treat bladder cancer. However, a significant drawback of these drugs is that they indiscriminately kill cells, often causing damage to the normal bladder mucosa or inflammation. Additionally, continuous dilution by urine or direct excretion leads to reduced effective drug concentrations, resulting in insufficient antitumor efficacy. Moreover, drugs struggle to penetrate the protective layer of basal cells lining the bladder, making curative effects in deep‐seated bladder tumors difficult to achieve. Finally, most antitumor drugs have significant side effects, potentially harming patients. Therefore, traditional bladder instillation therapy is often associated with high tumor‐recurrence rates. In a randomized clinical trial, among 201 patients with NMIBC who underwent TURBT and received intermittent gemcitabine intravesical instillations, tumor recurrence rates remained as high as 34% at 4 years post‐surgery [21]. Chen et al. reported that, among 127 patients with NMIBC who underwent TURBT, the 5‐year bladder‐tumor recurrence rates were 37.9% after doxorubicin treatment and 33.9% after BCG treatment [22]. Considering the limitations of traditional bladder instillation, several researchers have made improvements in recent years. For instance, some researchers have employed polymeric microspheres encapsulating paclitaxel for bladder instillation [23]. Compared with conventional paclitaxel instillation, these composite microspheres demonstrated sustained drug release and prolonged antitumor activity. Other researchers synthesized composite microspheres combining magnetite colloidal nanoparticles with Fe₃O₄@ALA‐Zn II, achieving significant anti‐bladder‐tumor effects through photosensitization and zinc‐ion activity [24]. Karavana et al. developed composite microspheres encapsulating antitumor drugs and hydrogels to enhance adhesion to bladder mucosa, thereby achieving sustained drug control [16]. However, prolonged hydrogel action may induce urinary tract irritation as a side effect. Xiong et al. used Pluronic F‐127–based microspheres to deliver the TGF‐β1 inhibitors LY3200882 and gemcitabine, leveraging synergistic effects to enhance anti‐bladder‐tumor activity while inhibiting tumor‐cell metastasis [25]. Although significant progress has been made in current research, the shortcomings of traditional bladder instillation have only been partially addressed. The novelty of PCTM microspheres lies in their pioneering integration of four key functions—tissue adhesion, tumor targeting, cellular penetration, and antitumor therapy—into a single biodegradable microsphere platform. This integration transcends simple functional stacking to temporally and spatially coordinated therapy. The first function is synergistic adhesion: TA initially adheres to the bladder mucosa, whereas the positively charged PTCM microspheres exhibit significant electrostatic attraction to the negatively charged bladder‐tumor regions, laying the foundation for sustained controlled drug release. The second mechanism involves synergistic targeting and aggregation: positively charged PTCM microspheres aggregate at negatively charged bladder‐tumor sites. Concurrently, R11 exhibits pronounced active targeting of bladder tumors; this synergy ensures microsphere enrichment at the tumor site following adhesion. Following adhesion‐controlled release, the membrane‐penetrating peptide R11 and the M3‐2K–synthesized RM peptide readily traverse tumor‐cell membranes and penetrate deeply into the tumors, creating favorable conditions for eradicating deep‐seated tumors. M3‐2K, an immunomodulatory peptide, exhibits excellent cellular uptake [13]. It disrupts mitochondrial function in tumor cells, induces cell‐cycle arrest, and ultimately triggers apoptosis. These multifunctional PCTM microspheres demonstrate effective tumor targeting, prolonged controlled drug release, exceptionally high cellular uptake efficiency, and rapid induction of tumor cell apoptosis, thereby achieving synergistic therapeutic effects.

3. Conclusion

We developed multifunctional microspheres for intravesical perfusion to eradicate bladder cancer by blocking the progression of the G1/S phase of the tumor cell cycle. The PCTM microspheres significantly enhance targeting, adhesion, and penetration into bladder tumor cells, facilitating intracellular delivery to bladder cancer cells and blocking DNA replication and cell‐cycle progression by downregulating the expression of HCFC1, CDC42, and cyclin A1 cell‐cycle proteins, ultimately leading to apoptosis. Both the in vivo and in vitro experimental results confirm that the PCTM microspheres possess potent antitumor properties and reliable biosafety. Therefore, the PCTM microspheres are a promising radical treatment for bladder cancer via bladder perfusion, and the data provided here demonstrate their potential for clinical applications.

4. Experimental Section

4.1. Materials

R11 and M3‐2K peptides were purchased from the Wuhan Branch of Hubei Qiangyao Biotechnology (Hubei, China). Common Cy Dye, PLA, DiD, MTT Assay Kit, PI Staining Solution, Ham's F‐12K medium, Annexin V‐FITC Apoptosis Detection Kit, Cell Counting Kit‐8 (CCK‐8), Calcein‐AM/PI Cell Viability/Cytotoxicity Assay Kit, human bladder cancer cell lines T24 and T5637, and normal urothelial cells (SVHUC‐1) were purchased from Shanghai Nuo He Biotechnology (Shanghai, China). BALB/c nude mice were purchased from Suzhou Hengxin Chen Biomedical (Suzhou, China). Hematoxylin staining solution and eosin staining solution, Anti‐Caspase‐3 Rabbit pAb and Anti‐Ki67 Rabbit pAb, Anti‐alpha smooth muscle Actin Rabbit pAb, β‐Actin Mouse mAb, Anti‐HCFC1 Rabbit pAb, Anti‐Cyclin A1 Rabbit pAb, Anti‐CDC42 Rabbit mAb, horseradish peroxidase–labeled goat anti‐mouse IgG, and RP‐labeled goat anti‐rabbit IgG were purchased from Wuhan Xavier Biotechnology (Wuhan, China).

4.1.1. Synthesis of RM Polypeptide

Briefly, Fmoc solid‐phase synthesis [26] was used to prepare double‐stranded polypeptides containing functional groups in a stepwise manner, and click chemistry (azide–alkyne cycloaddition reaction) was used to synthesize the RM composite protein by linking fragment I (containing azide radicals) with fragment II (containing dibenzo‐cyclooctyne) to form a stabilized triazole bond via a specific cycloaddition.

4.1.2. Synthesis of PCTM Multifunctional Microspheres

4.1.2.1. Aqueous‐Phase Solution Formulation (H₂O + PVA)

Polyvinyl alcohol (PVA) was dissolved in deionized water (concentration 1%–5% w/v), magnetically stirred (500 rpm, 50°C, 2 h) until complete dissolution, and then filtered through a 0.22 µm membrane and set aside for use. PVA was used as the stabilizing agent to prevent nanoparticle aggregation. Organic‐phase solution formulation (DCM + PLA + Cy5): PLA was dissolved in dichloromethane (DCM; 2%–5% w/v), the hydrophobic NIR fluorescent dye Cy5 (0.1%–0.5% w/w PLA) was added, and ultrasonication (40 kHz, 10 min) was performed to ensure uniform dispersion. PLA was used as the hydrophobic carrier to encapsulate Cy5 for fluorescence tracing.

4.1.2.2. Microfluidic Preparation of Microspheres (PLA + Cy5, PC)

The aqueous and organic phases were injected at a constant flow rate (3 mL min⁻¹ for the aqueous phase and 1 mL min⁻¹ for the organic phase) using a dual‐channel microfluidic technique, and the self‐assembly of the microspheres was induced by shear force. After collecting the emulsion, DCM was removed by spinning under reduced pressure for 2 h, and the PLA‐Cy5 microspheres were collected by centrifugation (12,000 rpm, 20 min) and washed three times with PBS.

4.1.2.3. TA Surface Modification (PLA + Cy5 + Ta, PCT)

The PC microspheres were dispersed in PBS (pH 7.4), and TA (1 mg mL⁻¹) was added and stirred at room temperature (300 rpm, 1 h).

4.1.2.4. Peptide Functionalization Modification (PLA + Cy5 + Ta + RM, PCTM)

The RM peptide was added to the PCT microsphere suspension at a molar ratio of 1:1 (RM:PCT) and incubated at 4°C for 12 h. Unbound peptides were removed by dialysis (MWCO 50 kDa, 24 h) and freeze dried.

4.1.2.5. Cell Passage and Culture

SV‐HUC‐1, T24, and T5637 cells: After the medium was discarded, the adherent cells were gently washed once with 2 mL of PBS to remove residual serum; 1 mL of 0.25% w/v trypsin‐EDTA solution was added, and the solution was incubated for 4 min, 30 s at 37°C to digest the cells. After microscopic confirmation of cell‐gap enlargement, the cells were pipetted and blown 15 times to induce complete detachment of the cells from the surface of the dish. The cell suspension was transferred to a 15 mL centrifuge tube and mixed with an equal volume of complete medium containing 10% fetal bovine serum to terminate the digestion, then centrifuged at 3000 rpm for 3 min. The supernatant was discarded, and the cell suspension was incubated at a 1:2 ratio with 0.25% w/v trypsin‐EDTA solution. The cell suspension was incubated at 37°C under 5% CO₂, and cell‐wall attachment and proliferation were analyzed after 24 h.

4.1.2.6. SEM and AFM Analyses

A Sigma 300 scanning electron microscope (ZEISS, Oberkochen, Germany) with a shooting voltage of 3 kV was used for the test model, and an Xplore 30 (Oxford) scanning electron microscope was used for the spectroscopy model. An appropriate amount of the powder was directly adhered to the conductive adhesive, sprayed with gold, and mounted on a Dimension Icon atomic force microscope (Bruker) to analyze the morphology of the microspheres and measure their diameters. A portion of each sample was ultrasonicated in ethanol solution, then a few drops of the well‐dispersed liquid were added dropwise on a mica sheet and scanned in the area of 20 × 20 µm to observe the morphology and roughness of each group of microspheres.

4.1.2.7. Surface Potential Analysis of Synthetic Microspheres

The sample was dispersed in 0.1% deionized water and ultrasonicated for 3 min. The distribution and zeta potential were measured using a nanoparticle size analyzer, and the average value was obtained from three tests.

4.1.2.8. HPLC–MS

Synthetic proteins were dissolved in a water/acetonitrile mixture containing 0.1% formic acid (70:30, v/v) and filtered through a 0.22 µm membrane. Then, 5 µL of proteins were injected to optimize the chromatographic conditions, and full‐scan and data‐dependent MS/MS spectra were acquired using an Orbitrap analyzer.

4.1.2.9. IR Spectroscopy

A small amount of the sample and an appropriate amount of dry KBr powder was ground thoroughly and pressed into a transparent tablet on a tablet press. The IR spectra of the background and sample were collected.

4.1.2.10. XPS Analysis

An appropriate amount of the sample was pressed and placed in the sample chamber of a Thermo Scientific K‐Alpha X‐ray photoelectron spectrometer. When the pressure was less than 2.0 × 10⁻⁷ mbar, the spot size was 400 µm, the operating voltage was 12 kV, and the filament current was 6 mA, the samples were fed into the analysis chamber. The full‐spectrum scanning fluence energy was 150 eV in 1 eV steps, and the narrow‐spectrum scanning fluence energy was 50 eV in 0.1 eV steps.

4.1.2.11. Kinetic and Molecular Docking Analysis

3D structures of the A chain (RM), D chain (2‐Azido+DBCO‐Mal), B chain (5‐FITC‐(Acp)‐(DOPA)‐G‐(DOPA)‐C‐(DOPA)‐G‐(DOPA)), and ADB holopeptides were constructed using PyMOL software [27]. Gaussian16 [28] software was optimized to generate RESP charges using Multiwfn [29]. Gromacs+ simulation software was used [30]. The force field applied was aber14sb_parmbsc1.ff, the water molecule was modeled as TIP3P, and Sobtop used the GAFF force field to generate the topology files for the B and D chains. After constructing the initial simulation system with the A, B, and D chains and ADB holopeptide, the truncation radius of 12 Å was calculated for the coulombic and van der Waals interactions, hydrogen bonding was predicted using the LINCS algorithm, and the simulation results were visualized using the Gromacs and PyMOL programs. Secondary structure calculations were performed using DSSP (Define Secondary Structure of the Protein) software. The program HDOCK [31] was used for protein–protein and protein–DNA/RNA docking, and docking scores were calculated based on the ITScorePP or ITScorePR iterative scoring functions. A more negative docking score indicated greater binding potential and stronger interactions in the binding model. Considering that the docking scores of protein–protein complexes in the PDB are usually approximately –200 or better, we empirically defined a docking confidence score to indicate the binding likelihood of two molecules as follows: confidence_score = 1.0/[1.0+e0.02*(Docking_Score+150)]. When the confidence score was above 0.7, the two molecules were highly likely to bind; when the confidence score was between 0.5 and 0.7, the two molecules were likely to bind; and when the confidence score was below 0.5, they were unlikely to bind.

4.1.2.12. Molecularization

Owing to the large number of atoms, geometry optimization was performed using the GFN2‐xTB method in XTB 6.6.1 [32]. Furthermore, singlet‐point‐energy calculations were performed using the Gaussian D01 software package at the B3LYP(D3)/6‐31G(d) level of theory to obtain a stable wave function. The noncovalent interactions (NCl) and electrostatic potential were determined using the Multiwfn 3.8 (dev) program [33].

4.1.2.13. Loading of RM Peptides into PTCM Microspheres

First, 1.00 ± 0.02 mg of lyophilized PTCM microspheres (three biological replicates) were precisely weighed. Then, the microspheres were immersed in 10 mL of PBS solution preequilibrated at 37°C containing 0.1 mg mL⁻¹ RM peptide. The samples were incubated at 80 rpm in a constant‐temperature shaker for 12 h and then centrifuged at 10,000 rpm for 5 min to separate the supernatant. Detection was performed using an HPLC autosampler, with three replicate injections per sample. The loading capacity was calculated using a standard curve. Data were processed using GraphPad Prism 9.0 (GraphPad Software, California, USA).

4.1.2.14. Release of RM Peptides Loaded into the PTCM Microspheres

Microspheres loaded with 1.00 ± 0.02 mg were precisely weighed, and three biological replicates were suspended in 10 mL of PBS. The release was conducted at 37°C in a constant‐temperature shaker at 80 rpm. The sampling time points included the initial 12 h, followed by 1, 3, 5, 7, 9, 11, 14, and 20 days. After each 1 mL sample collection, an equal volume of fresh PBS prewarmed to 37°C was immediately replenished. Automatic HPLC injection detection was performed, and each sample was analyzed in triplicate. The concentration of the RM peptide was determined using HPLC, enabling the calculation of cumulative release amounts and release profiles.

4.1.2.15. NIR Thermal Imaging

Four groups of microspheres (P (PLA), PC, PCT, and PCTM) were used in the blank, T24‐cell, and nude‐mouse groups. A background signal was collected from the blank group for subsequent background deduction from the images, 3 mL of the four microspheres at a concentration of 5 µg mL⁻¹ were incubated with T24 cells for 4 h (37°C, 5% CO₂), and 2 mL of the four microspheres at a concentration of 5 µg mL⁻¹ were injected into the bladders of nude mice through a 24G intravenous needle and subsequently imaged separately using a NIR imager.

4.1.2.16. Cell Counting Kit‐8

After trypsin digestion of the T24 cells, a single cell suspension was made, and 100 µL of the cell suspension with a count of 1 × 10⁴ was added to each well of a 96‐well plate. The cells were incubated at 37°C for 24 h in a 5% CO₂ incubator. Afterward, 100 µL of PCTM medium containing 100, 50, 20, 10, 5, 2.5, and 1.25 µg mL⁻¹ was added, and three wells were set up for each concentration. The negative control group was treated only with PBS. The plates were incubated for 48 h, and 10 µL of CCK‐8 solution was added to each well 2 h before the end of the incubation period. The OD value of each well was determined using an enzyme marker at 450 nm and was calculated as follows: cell viability (%) = (OD drug group—OD blank group)/(OD negative control group—OD blank group) × 100. CCK‐8 assays of SV‐HUC‐1 and T5637 cells were performed in the same manner.

4.1.2.17. Live–Dead Cell Assay

After the T24 cells were recovered, the cells were digested with 1 mL of 0.25% (w/v) trypsin‐EDTA solution, the quantity was adjusted to 5 × 10⁴ cells mL⁻¹, and 100 µL of cells per well were inoculated into a 96‐well plate. Then, 100 µL (5 µg mL⁻¹) of the P, PC, PCT, and PCTM microspheres were added to three wells of each group. Calcein‐AM (100 µL, 2 µm) and PI (4.5 µm) were added to each well and incubated for 30 min, and the live (yellow–green fluorescence) and dead (red fluorescence) cells were detected using a fluorescence microscope equipped with an excitation filter at 490 ± 10 nm. Live and dead SV‐HUC‐1 and T5637 cells were also analyzed.

4.1.2.18. Targeting Assay

In order to verify the function of the RM peptide, we used 1 µg/mL of RM peptide, which is equivalent to the total amount of RM peptide loaded in 5 µg/mL PCTM microspheres. After trypsin‐induced digestion of DiI‐labeled T24 and T5637 cells and DiD‐labeled SV‐HUC‐1 cells, T24 and T5637 cells with a cell count of 1 × 10⁵ were co‐cultured with SV‐HUC‐1 cells. After cell attachment, 1 µg mL⁻¹ of the RM peptide solution was added to 12 wells at 12, 6, and 3 h according to the “countdown method.” The culture solution was discarded, and cells were washed and fixed with paraformaldehyde (0.5 mL). The fluorescence intensity of each well was observed using an inverted fluorescence microscope and semiquantitatively analyzed using ImageJ software.

4.1.2.19. Membrane Penetration Assay

T24, T5637, and SV‐HUC‐1 cells were digested and centrifuged, and 2.5 × 10⁵ cells were counted in each group and co‐cultured in 24‐well plates. The countdown method was used to add 1 µg mL⁻¹ of RM peptide into the wells of the plates at 12, 9, 6, and 3 h, followed by 400 µL of DID staining for 15 min and paraformaldehyde fixation for 15 min in each well. The fluorescence intensity of each well was analyzed using an inverted fluorescence microscope and semiquantitatively analyzed using ImageJ software.

4.1.2.20. Cell Scratch Assay

After trypsin digestion of the T24 and T5637 cells, 5 × 10⁵ cells were seeded into each well of a plate. Then, a 20 µL pipette tip was used to make a cell scratch along a straight edge, and the old medium was sucked out to leave a cell gap. Microscopic photographs were taken as a 0 h control, then medium containing 5 µg mL⁻¹ PCTM was added to each group of cells. The width of the scratches was observed microscopically at 6, 12, and 24 h, and the cells were analyzed semiquantitatively using ImageJ.

4.1.2.21. Clone Formation Assay

T24, T5637, and SV‐HUC‐1 cells were digested with trypsin to obtain single‐cell suspensions. A total of 700 cells were counted and inoculated in each well, and then P, PC, PCT, and PCTM medium containing 5 µg mL⁻¹ were added. After culturing for 10 days, the cells were fixed with paraformaldehyde for 60 min, stained with crystal violet for 20 min, and photographed using a digital camera.

4.1.2.22. Flow Cytometry Apoptosis Assay

SV‐HUC‐1, T24, and T5637 cells were inoculated into 6‐well plates, and 1 × 10⁶ cells were counted in each well. The cells were treated with a 10 µg mL⁻¹ PCTM microsphere solution, and the single‐cell suspension was collected after washing with PBS and incubating with Annexin V and PI according to the kit's instructions. Finally, the apoptosis rate was analyzed using flow cytometry at 0, 3, 6, and 12 h.

4.1.2.23. Flow Cytometry Cycle Assay

Trypsin solution was used to digest the SV‐HUC‐1, T24, and T5637 cells, which were then inoculated into 6‐well plates, and the cells (1 × 10⁶) were counted in each well. Each group of cells was treated with 10 µg mL⁻¹ of the PCTM microsphere drug, washed with PBS, and fixed with 70% ethanol, followed by collection of the single‐cell suspension. The cells were stained according to the Annexin V/PI kit instructions, and the cell cycle was analyzed using flow cytometry at 3, 6, 9, and 12 h.

4.1.2.24. Western Blot Analysis

T24 cells were inoculated in a 6‐well plate, and fresh media containing PCTM microspheres at concentrations of 0, 2, 4, and 8 µg mL⁻¹ were added. After processing for 24 h, RIPA buffer was added and cleaved for 30 min, and the sample was subjected to electrophoresis and membrane transfer. The primary antibodies were HCFC1 (1:1000), actin (1:5000), cyclin A1 (1:1000), and CDC42 (1:2000). The secondary antibody was horseradish peroxidase–labeled anti‐rabbit/mouse IgG (1:5000). Finally, the expression differences between the HCFC1 protein, cyclin A1, CDC42 protein, and the internal reference (actin) were compared.

4.1.2.25. Transcriptome Sequencing

T24 and T5637 cells were inoculated in a 6‐well plate, and fresh medium containing 5 µg mL⁻¹ of PCTM microspheres was added to the experimental group. Only the medium was added to the control group. After 24 h of treatment, TRIzol reagent was used to cleave the cells and RNA was extracted via centrifugation. The three experimental groups and three control groups were sent to Shanghai Ouyi Biomedical Technology for transcriptome sequencing.

4.1.2.26. In Situ Bladder Transplantation Tumor Model

Male BALB/c naked mice weighing 18–22 g at 6–8 weeks of age were intraperitoneally injected with 1% sodium pentobarbital after fasting for 12 h. The perineum and urethral opening were disinfected with 75% ethanol, and a 24 g injection needle was inserted into the urethra until the urine flowed out naturally. The needle core was rotated to establish a model of mechanical damage to the bladder mucosa, and 100 µL of the T24 cell suspension containing 1 × 10⁷ mL⁻¹ was injected via an indwelling needle. On the seventh day after the operation, an animal C‐arm X‐ray machine was used for bladder contrast confirmation, and the bladder tissues of two naked mice were randomly cut for H&E staining and immunohistochemical testing (Ki‐67, caspase‐3).

4.1.2.27. Subcutaneous Transplantation Tumor Model

After disinfection, a 1 mL syringe was used to puncture the skin of the right posterior axillary line of naked mice at an angle of 30° with the subcutaneous tissue, and 200 µL (5 × 10⁶ mL⁻¹) of T24 suspension was slowly injected. A vernier caliper was used to measure the 3D length of the tumor every day, and the volume of the tumor was calculated according to the ellipsoid formula: V = π/6 × length × width × height. When the average tumor volume reached 100 mm³, H&E staining was performed to confirm and initiate drug intervention. All animal experiments were conducted at the Experimental Animal Center of the Chaohu Hospital of Anhui Medical University and were approved by the Experimental Animal Ethics Committee of Anhui Medical University (grant number KYXM‐202309‐015).

4.1.2.28. In Vivo Bladder‐Adhesion Experiment

Balb/c‐nude mice were anesthetized with sodium pentobarbital, and then 2 mL of PC, PCT, and PCTM microspheres (5 µg mL⁻¹) were injected into the bladders of naked mice through an indwelling needle. The mice were sacrificed at 0.5, 2, 6, 12, and 24 h, after which the bladders were removed and the tissues sequestered. The fluorescence area and intensity of the bladder samples were analyzed using an inverted fluorescence microscope, and semiquantitative analysis was performed using ImageJ.

4.1.2.29. In Vivo Bladder‐Penetration Experiment

After Balb/c‐nude anesthesia was stabilized, 2 mL (5 µg mL⁻¹) of the PC, PCT, and PCTM microspheres were instilled into the bladder through an indwelling needle. The mice were euthanized at 0.5, 2, 6, 12, and 24 h; their bladders were removed; and full‐layer slices of the bladders were prepared. A fluorescence microscope was used to observe bladder wall penetration of the three groups of microspheres; ImageJ was used for semiquantitative analysis.

4.1.2.30. Tumor‐Suppression Experiment

In situ bladder‐tumor naked‐mouse model: Twenty naked mice with bladder cancer were randomly divided into PBS, P, PC, PCT, and PCTM groups. Each group was perfused with 2 mL (5 µg mL⁻¹) of the microsphere suspension into the bladder, once a week for 4 consecutive weeks. Weight changes in the animals were measured weekly, and the survival rate of naked mice with bladder cancer in situ was determined. Four weeks later, the mice were euthanized; the heart, liver, spleen, lungs, and kidneys were removed; and blood was collected for histopathological and biochemical analyses. Finally, the bladders of the naked mice were removed to compare changes in bladder tumor size in the treatment group.

4.1.2.31. Tumor‐Suppression Experiment of the Subcutaneous‐Tumor Naked‐Mouse Model

Successfully modeled subcutaneous‐tumor naked mouse 1 was randomly divided into PBS, P, PC, and PCT groups (three groups), and a PCTM group (four groups). A fixed subcutaneous injection at a concentration of 5 µg mL⁻¹ and 2 mL into the right armpit was administered every week for 4 weeks of continuous treatment. Weight changes and survival rates of the naked mice were evaluated weekly. After 4 weeks, the mice were sacrificed, and the heart, liver, spleen, lungs, kidneys, and blood were collected for histopathological and blood biochemistry tests. Finally, the bladders of the naked mice were removed to compare the changes in bladder tumor size in each group.

4.1.2.32. Statistical Analysis

Measurements were expressed as mean ± standard deviation, and comparisons between two groups were made using Student's t‐test. Three or more groups were compared using a one‐way analysis of variance (ANOVA). Graphs were plotted and statistically analyzed using GraphPad Prism 9.0 (GraphPad Software, USA) and Origin 2021 (OriginLab, Northhampton, MA, USA); ChemDraw 22.0 (PerkinElmer, Boston, MA, USA) was used to write the molecular chemical formulas; Avantage 5.9 (Thermo Fisher Scientific, USA) was used to analyze the XPS data; FlowJo 10.8 (BD, USA) was used for flow cytometry analysis; R4.3 (New Zealand) was used to analyze and graph the sequencing‐related data; ZEISS ZEN 3.8 (Carl Zeiss, Germany) and ZEISS ZEN 3.8 (Carl Zeiss, Germany) were used for graphing and statistical analysis; and Image J‐win64 was used for fluorescence photo data analysis. Differences were considered statistically significant at p < 0.05.

Author Contributions

SW.Y. and Y.H. curated and investigated the data. Y.Q. was responsible for conceptualization and methodology. C.L. supervised and validated the study. Z.L. was responsible for funding acquisition, formal analysis, editing, and manuscript writing.

Conflicts of Interest

The authors declare no conflict of interest.

Supporting information

Supporting File 1: adhm70648‐sup‐0001‐SuppMat.docx.

Supporting File 2: adhm70648‐sup‐0002‐TableS1.xls.

Data Availability Statement

The data that support the findings of this study are available on request from the corresponding author. The data are not publicly available due to privacy or ethical restrictions.