A live biohybrid bacterial therapy based on engineered Serratia marcescens

Domain: neuroscience
Published: 2026-04-07
Editorial outcome: major_revision

Abstract
Bacterial therapeutics hold great promise for cancer treatment by targeting oxygen-poor tumor regions and complementing existing therapies. However, current approaches often struggle with safety concerns and complex engineering. Developing a safe, effective delivery platform relying entirely on natural bacterial biosynthesis remains a challenge. Here we show that attenuated Serratia marcescens serves as a powerful biohybrid platform for cancer therapy by leveraging its natural biosynthesis of prodigiosin, a photosensitive pigment. We engineer S. marcescens to yield high prodigiosin levels, which exhibit strong intrinsic anti-cancer activity and near-infrared photosensitivity. In female mouse models of melanoma and colorectal cancer, this platform triggers robust systemic immune responses, including enhanced T cell recruitment and long-term memory against tumor recurrence. Furthermore, the bacteria induces tumor cell death via mitophagy, while photothermal properties of prodigiosin enables rapid, light-controlled bacterial clearance post-treatment. These findings establish S. marcescens as a versatile, self-regulating biosynthetic platform for precise and safe cancer immunotherapy.

Introduction
Bacteria-based delivery systems have garnered attention due to their unique capabilities in targeting and treating various diseases, including cancer and bacterial infections1-4. These systems leverage the natural properties of bacteria-such as hypoxia-targeting capability, versatile biosynthetic machinery, and immunostimulatory potential-to address drug resistance5, 6. Various bacterial strains have been employed as delivery vehicles to date, including Salmonella typhimurium VNP200097-10, Clostridium novyi-NT11-14, Escherichia coli15, 16, Listeria monocytogenes17-19 and so on. Among these, VNP20009 and C. novyi-NT have undergone attenuation to improve safety profiles and have reached Phase I clinical trials20-22, whereas others remain in preclinical development. Conventional bacterial therapies predominantly inhibit tumor growth indirectly by activating the immune system23, 24. However, their direct ability to kill tumor cells remains limited and often necessitates intricate physical or chemical modifications to facilitate drug or therapeutic media delivery—a process that is time-intensive, inefficient, with limitations in drug release efficiency and scalability.

To address this limitation, researchers have employed genetic engineering to amplify the innate anti-tumor immune function, thereby circumventing the reliance on external interventions. For instance, Andrew et al2. engineered Escherichia coli Nissle 1917 to deliver tumor-specific neoantigenic epitopes, thereby activating specific CD4+/CD8+ T cells and significantly enhancing the efficacy of antitumor immunotherapy. Similarly, Wang et al25. modified Salmonella typhimurium VNP20009 to secrete granulocyte-macrophage colony-stimulating factor (GM-CSF), which persistently induced the reprogramming of M2-type tumor-associated macrophages into M1-type anti-tumor macrophages, thereby boosting the anti-tumor immune response. While these strategies effectively inhibit tumor growth by heightening bacterial immune activation, their direct tumor-cell-killing capabilities remain constrained. Additionally, developing a safe, biosynthesis-based platform for anticancer drug delivery continues to pose significant challenges.

Bacteria serve as an important platform for anticancer natural products, such as polyketide/macrolide antibiotics and non-ribosomal peptides26, 27. However, their complex synthetic pathways and difficulties in heterologous expression often constrain production yields28-30. By contrast, microbial pigments with antitumor abilities can typically be produced in large quantities, making them an attractive alternative31. Among these is Serratia marcescens–derived prodigiosin, a linear tripyrrole bright red natural product with demonstrated antibacterial, antioxidant, and anti-cancer properties32-34. Prodigiosin suppresses the Wnt/β-catenin pathway by downregulating the phosphorylation of key regulatory factors, thereby reducing β-catenin activation, lowering cyclin D1 expression, and ultimately inducing tumor cell apoptosis to inhibit tumor progression.35. However, its application in cancer therapy has been hampered by challenges including its low bioavailability, chemical instability in varying environments, and short systemic half-life36. Unlocking its potential requires a deeper understanding of its properties and innovative mechanisms for its application.

In this study, we report a biohybrid strategy based on attenuated S. marcescens for cancer therapy that leverages the natural biosynthesis of photosensitive prodigiosin. This biohybrid approach addresses both safety and efficacy concerns: it delivers potent antitumor effects in vitro and in vivo inducing robust cell death, mainly through mitophagy-driven necroptosis and caspase-3-independent apoptosis in tumor cells, promotes immunological responses (dendritic cell maturation, T cell recruitment, and macrophage polarization), and rapid bacterial clearance via prodigiosin-mediated photothermal ablation. Collectively, these findings indicate that the engineered S. marcescens can serve as a robust chassis for delivering photosensitive natural products, offering a valuable avenue for more precise and effective bacterial cancer therapies (Figure 1).

Results
Discovery of Prodigiosin Exhibiting Near-Infrared Fluorescence and Photothermal Effects.
Starting from S. marcescens NRRLTB-1481, we developed an engineered strain, S. marcescens JC11 (hereafter referred to as SMM), through iterative UV irradiation and visual screening for intensified red pigmentation, indicating enhanced prodigiosin biosynthesis (Supplementary Fig1, 2). Random mutagenesis yielded SMM, capable of producing a competitive 8.3±0.5 g/L prodigiosin (upper range for native S. marcescens under optimized conditions) in a 5-L bioreactor within 48 h (Supplementary Table 2: representative production studies). While yields of up to 10.25 g/L have been achieved through techniques like promoter engineering of OmpR and PsrA37, SMM offers a scalable and cost-effective alternative, particularly valuable for downstream translational and clinical applications.

Interestingly, this high-yield phenotype is tightly regulated by temperature; consistent with the known regulatory network, prodigiosin biosynthesis peaks at 30℃ because the LysR-family repressor HexS silences the pigA-N promoter at ≥37℃38, 39. Activators such as PigP remain expressed at higher temperature but cannot override HexS-dependent repression40. A serendipitous live-cell imaging experiment revealed that prodigiosin emits fluorescence under 808 nm excitation (Figure 2a). Emission scanning identified a primary peak at approximately 940–950 nm (near-infrared I), extending to around 1300 nm (near-infrared II), with a first-order fluorescence lifetime of 1208 ns (Figure 2 c,d). At a physiological temperature of 37 degrees, the bacteria undergo physiological degradation, with a clearance efficiency of over 80% observed within seven days (Supplementary Fig3 a,b). This characteristic of the strain also indicates its potential to develop into an excellent platform for integrated diagnosis and treatment delivery.

A 5 mM solution of purified prodigiosin in ethanol exhibited potent photothermal conversion, with temperatures rapidly exceeding 60℃ (under 4.5 W/cm² irradiation) (Figure 2e,f; Supplementary Fig4a). Importantly, the bacteria with pre-synthesized prodigiosin, acting as living photothermal agents also demonstrated this capability (Supplementary Fig5a). Aqueous suspensions of SMM showed a rapid, concentration-dependent temperature increase under 808 nm laser irradiation (4.5 W/cm²), with a concentration of 1×10¹¹ CFU/mL reaching over 60℃ in just 3 minutes (Figure 2 g,h; Supplementary Fig5). This bacterial agent with pre-synthesized prodigiosin also exhibited excellent photostability across multiple heating cycles (Supplementary Fig5b).This temperature threshold (~60℃) was validated as being sufficient for thermal ablation; a 3-minute treatment at 60℃ resulted in a greater than 8-log reduction in bacterial viability, effectively sterilizing the culture (Supplementary Fig6, S7). However, routine purification and direct use of prodigiosin are hampered by the co-production of proteases and surfactants, as well as prodigiosin’s structural complexity and moderate instability. Building on the hypoxia-targeting capability of SMM and the intrinsic anticancer potential of biosynthesized prodigiosin, we hypothesize that SMM can serve as a cellular drug‑delivery host, delivering prodigiosin specifically to tumor sites and enabling a living theranostic strategy. To establish a robust, dual-layered safety profile for this approach, we first engineered an attenuated strain based on SMM, by deleting the msbB gene (hereafter referred to as SM), which is critical for the biosynthesis of endotoxic lipid A; the successful knockout was confirmed by PCR of genome (Figure 2i). Layered on top of this genetic modification, the photothermal property of the bacteria-produced prodigiosin permits on-demand bacterial ablation via 808nm near-infrared irradiation, substantially enhancing the overall and controllable safety of this living therapeutic.

Biohybrid Strategies to Attenuate Serratia marcescens and Achieve Targeted Tumor Localization
Building upon the enhanced safety profile of the genetically detoxified SM strain, we introduced multi-functional microencapsulation strategy using a polymeric material (Figure 3a). This encapsulation serves two key purposes: first, it minimizes systemic exposure and reduces potential immunogenicity by limiting bacterial surface antigen presentation. Second, the polymeric shell provides abundant surface carboxyl groups, enabling efficient amide coupling of targeting peptides and minimizing off-target delivery. Notably, even before msbB deletion, SMM does not cause hemolysis on blood agar plates, thus further highlighting its reduced virulence and improved biosafety (Figure 3b). Like Escherichia coli Nissle1917 (ECN) and Salmonella typhimurium VNP20009 (VNP20009), SM is a facultative anaerobe; however, under strict anaerobic conditions at 37℃, SM growth is significantly inhibited (Supplementary Fig8). Interestingly, the degree of anaerobic growth inhibition observed for SM was less pronounced than that of ECN, and roughly equivalent to VNP20009 (Supplementary Fig8). Deletion of msbB (SM) lowered lipopolysaccharide levels by 39.8% relative to the wild-type strain, as confirmed using Limulus amebocyte lysate assays (Figure 3c). In a murine model, SMM, ECN, and JM109 all exhibited similar high mortality ( >50% by day 4) following 1×108 CFU/mice intravenous administration (Figure 3d). Strikingly, SM showed 100% survival, demonstrating elimination of SMM lethality upon msbB deletion and confirming its critical role in virulence (Figure 3d).

A separate chemical surface engineering platform was then established to improve tumor selectivity (Figure 3a). Polydopamine (PDA), carboxymethyl cellulose (CMC), and carboxylated chitosan (CMCS) were initially tested as coating materials (Supplementary Fig9a–d). PDA was excluded due to potential fluorescence interference, and CMCS posed handling challenges arising from high viscosity. Consequently, CMC was selected for encapsulation (Supplementary Fig9a–d). Transmission electron microscopy confirmed a successful coating process, and the 808 nm-induced fluorescence remained intact (Supplementary Fig9a). The CMC coating achieved 98.2% efficiency without compromising cell viability (>90% viability; Supplementary Fig9). In parallel, the zeta potential decreased by 59.5%, while the particle size peak increased from 615.14 nm to 712.38 nm (Supplementary Fig9 b,c). Energy-dispersive X-ray spectroscopy (EDS) further validated Ca2+ doping and CMC functionalization (Supplementary Fig9d). Therefore, we selected CMC for encapsulation, naming the resulting composite SM@CMC, where the abundant carboxyl groups of CMC provide a rich resource for subsequent modification reactions.

To further enhance tumor targeting, we conjugated the cyclic peptide KP101-which exhibits high affinity for tumor-associated PD-L1 receptors41-onto SM@CMC via EDC/NHS coupling, yielding SM@CMC-KP101 biohybrids. FITC-labeled KP101 was employed for fluorescence microscopy and flow cytometry, confirming specific peptide–bacterium interactions (Figure 3e). Infrared spectroscopy revealed a new characteristic peak at approximately 1500 cm-1 in SM@CMC-KP101 (red arrow in Figure 3f), which is typically assigned to amide groups (e.g., amide II band) or peptide bond vibrations. This finding indicates that KP101 was successfully conjugated to the CMC-modified bacterial surface, providing direct evidence of amide bond formation and thereby validating the construction of SM@CMC-KP101 biohybrids. Following peptide grafting, the particle size increased to 955.4 nm, whereas the zeta potential decreased to −8.917 mV (Figure 3g, h). Flow cytometry analysis revealed a significant shift in FITC signal for SM@CMC-KP101, indicating successful binding of FITC-KP101(Figure 3i). The results of the transmission electron microscope (TEM) reveal that the bacterial surface, after being wrapped and modified with CMC and KP101, is rougher and shows distinct granular modifications, indicating the successful construction of our hybrid structure (Figure 3j). Furthermore, this biohybrid strategy did not affect bacterial viability, with the survival rate being essentially consistent with that of the unmodified strains (Supplementary Fig10). Quantitative analysis revealed that approximately 6 nmol of KP101 were anchored to every 108 colony-forming units (CFU), corresponding to ~3.6×107 peptides per bacterium6. The interaction pattern between bacteria and tumor cells involves the KP101 peptide on the bacterial surface targeting the PD-L1 receptor, which is highly expressed on the surface of tumor cells, allowing for selective binding (Figure 3k).

Synergistic in vitro Tumor Elimination and Immune Potentiation by SM@CMC-KP101
To evaluate the therapeutic potential of SM@CMC-KP101, we first examined its tumor-killing capacity and immunomodulatory effects in vitro. The intrinsic multi-wavelength fluorescence of prodigiosin (λmax = 592 nm, λex = 434 nm) in SM enabled real-time visualization of cellular uptake without additional labeling (Supplementary Fig11). Microscopic evaluation revealed that B16F10 and CT26 tumor cells internalized the bacteria through an endocytic mechanism (Figure 4a, b; Supplementary Fig12,13). CCK-8 assays using B16F10 and CT26 tumor cells demonstrated the cytotoxicity of SM@CMC-KP101 (IC50 < 4×107 CFU/mL, equivalent to 500 nmol PG) in a concentration-dependent manner (Figure 4c). Initial experiments using Annexin-V/PI staining in B16F10 murine melanoma cells showed an apparent increase in cell death following treatment with prodigiosin (PG) alone and SM@CMC-KP101 (Supplementary Fig14). However, given the spectral overlap between prodigiosin's emission (λmax = 592 nm, λex = 434 nm) and the emission spectra of Propidium Iodide (PI), Fluorescein isothiocyanate (FITC) channels, and particularly considering the FITC-like emission of Annexin-V, commonly used for apoptosis assays, these results are not conclusive evidence of apoptosis due to potential signal interference (Supplementary Fig14). Further mechanistic investigations are detailed in a subsequent section.

To determine the impact of surface modifications on bacteria cell entry, we directly compared the parental strain SMM, the attenuated mutant SM, the polymer-coated SM@CMC, and the KP101-functionalized SM@CMC-KP101 for their internalization efficiency in B16-F10 cells (Figure 4d). Using an amikacin protection assay, we incubated B16-F10 cells with equal inocula of each strain for 1 hour, then removed extracellular bacteria with 50 µg/mL amikacin42. After lysing the cells with 0.1% Triton X- 100, intracellular colony-forming units (CFU) were enumerated (Figure. 4d). The SM strain showed no significant difference in internalization compared to the parental SMM strain (both ≈0.09~0.10% internalization of the input dose). While CMC coating produced a slight increase, KP101 functionalization significantly enhanced bacterial entry, resulting in ≈0.19% internalization, representing a 107% improvement over SM under identical conditions (Figure. 4d). These results indicate that the enhanced cellular entry is primarily attributable to the KP101 peptide ligand rather than to msbB deletion or CMC coating.

Next, we assessed the immunostimulatory function of SM@CMC-KP101 in bone marrow–derived dendritic cells (BMDCs), which are pivotal antigen-presenting cells (APCs). In bone marrow-derived DC (BMDC) models, co-culture with SM@CMC-KP101 markedly enhanced the proportions of CD11c+CD80+CD86+ subsets (1.08fold increase vs. PBS treated group) (Figure 4e). Representative flow cytometric plots of bone marrow–derived dendritic cells (BMDCs) stained for CD11c+, CD80+ and CD86+ after the indicated treatments (Figure 4e). The data also indicated that SM-EV (23.83%), ECN (26.74%), and PG (20.82%) induced BMDC activation to varying degrees (Figure 4e). These findings emphasize the capacity of SM@CMC-KP101 to promote DC maturation and improve antigen-presenting activity. Reprogramming tumor-associated macrophages (TAMs) from anti-inflammatory M2 to pro-inflammatory M1 phenotype represents a critical strategy for reversing immunosuppressive microenvironments, we investigated macrophage polarization using Raw264.7 cells. SM@CMC-KP101 treatment significantly increased mRNA levels of M1 markers (TNFα, 62.94-fold; iNOS, 107.3-fold) with comparatively minimal changes in M2 markers (CD206, 3.3-fold; IL-10, 4.6-fold) (Figure 4f–i). Similar trends were observed in bone marrow–derived macrophages (BMDMs), where SM@CMC-KP101 upregulated M1-associated genes (iNOS, TNFα) while leaving M2 markers (CD206, IL-10) largely unaffected (Supplementary Fig15). Collectively, these data highlight the dual tumoricidal and immunostimulatory capabilities of SM@CMC-KP101 in vitro.

We developed a biohybrid drug delivery system loaded with prodigiosin for direct tumor cell killing. Furthermore, this system exhibits multifunctional capabilities (Figure 4j). Firstly, it significantly enhances dendritic cell (DC) maturation in vitro by increasing the proportion of CD11c+CD80+CD86+ subpopulations (1.8-fold increase compared to controls), thereby promoting antigen cross-presentation. Additionally, our system markedly increases the mRNA expression levels of M1 markers in M2-type macrophages (both the RAW264.7 line and primary BMDMs) by several hundred folds in vitro relative to untreated groups, while maintaining minimal changes in the immunosuppressive cytokine IL-10. This multimodal immunomodulatory activity—combining direct tumor cell killing with reprogramming of both innate and adaptive immunity—positions this platform as a promising bacterial-mediated therapeutic approach for solid tumor eradication. These encouraging in vitro results led us to investigate whether this treatment could subsequently stimulate adaptive immunity in vivo.

In vivo Evaluation of SM@CMC-KP101 in the CT26 Tumor Model
To ensure the safe and effective use of SM@CMC-KP101, we performed dose-escalation studies via intratumoral (i.t.) and intravenous (i.v.) administration in CT26 tumor-bearing mice (Supplementary Fig16a, b). This route-dependent approach determined the maximum tolerated dose of SM@CMC-KP101 to be 1×1010 CFU/mice (i.t.) and 1×108 CFU/mice (i.v.). Intratumoral injections used concentrations from 1×109 to 1×1013 CFU/mL (100 μL/mice, equivalent to 1×108-1×1012 CFU/mice), while intravenous injections ranged from 1×107 to 1×1011 CFU/mL (100 μL/mice, equivalent to 1×106-1×1010 CFU/mice) (Supplementary Fig16a-b). Based on these safety limits for SM@CMC-KP101, we conducted a 11-day therapeutic evaluation, monitoring tumor volume and body weight (Supplementary Fig16c-f). The optimal doses for antitumor activity without significant toxicity were 1×1010 CFU/mice (i.t.) and 1×108 CFU/mice (i.v.), leading to ≥97% tumor volume reduction.

To ensure the safety of the experiment, we performed intratumoral injections of ECN (1×1010 CFU/mice), VNP20009 (1×1010 CFU/mice), SM-EV (1×1010 CFU/mice), SM@CMC-KP101 (1×1010 CFU/mice) (Figure 5a). The injection area was illuminated with 4.5 w/cm² of 808 nm near-infrared light for 3 minutes to reach temperatures of 60℃ or higher. We also intravenously injected SM@CMC-KP101 (1×108 CFU/mice) in CT26 tumor-bearing mice. First, leveraging the inherent near-infrared (NIR) fluorescence emission of prodigiosin, we conducted in vivo imaging immediately following intratumoral administration of SM@CMC-KP101 (1×1010CFU/mice). Real-time observation revealed excellent in vivo imaging capabilities (Figure 5b), demonstrating stable biodistribution in major organs 24 hours post-injection. Notably, both heat-inactivated and live SM@CMC-KP101 exhibited strong tracer effects, highlighting the robust diagnostic potential of our live bacterial drug. However, due to the low intravenous injection dose (1×108 CFU/mice), we were unable to observe clear bacterial localization via in vivo imaging. Therefore, our current imaging data is presented exclusively for intratumoral injections.

In subsequent therapeutic studies, intratumoral injection of SM@CMC-KP101 followed by 808 nm photothermal therapy achieved effective intratumoral temperatures (≈60℃) while maintaining skin/normal-tissue temperatures at ≤43–45℃ (Supplementary Fig17). Following the treatment, normal organs showed no detectable bacteria at baseline; on day 1 (D1), they were either negative or exhibited only transient low signals (<102 CFU/g), and by D14, all organs achieved complete bacterial clearance (Figure 5c). Within tumors, the mean burden decreased from ≈ 1.3×109 CFU/g before treatment to≈4.25×103 CFU/g on D1, ≈5.5-log reduction, and all tumors were cleared by D14 (Figure 5c). Concurrently, the robust hyperthermia created three distinct phases of tumor destruction: (i) an acute phase (0-48 h) characterized by visible necrosis and hemorrhagic crust formation; (ii) a transition phase (3-7 d) marked by complete scab detachment with underlying granulation tissue; and (iii) a resolution phase (10-15 d) resulting in full epithelialization without scarring (Figure 5d). This near-complete bacterial clearance minimized infection risk while leaving behind bacterial debris that potentially activates local immune responses.

To elucidate the impact of SM@CMC-KP101 on the tumor immune microenvironment, we analyzed key immune cell populations. After 24 hours, dendritic cells (DCs) isolated from the tumor-draining lymph nodes exhibited enhanced maturation, with the proportion of CD11c+CD80+CD86+ DCs significantly increased in the SM@CMC-KP101(i.t.) (24.6%) and SM@CMC-KP101(i.v.) (16.8%) groups compared to the PBS treated group (9.9%) (Figure 5e, Supplementary Fig19a). Similarly, the proportion of CD3+CD8+ T cells was elevated in the SM@CMC-KP101(i.t.) (26.3%) and SM@CMC-KP101(i.v.) (32.9%) groups relative to controls (21.0%) (Figure 5f, Supplementary Fig19b). Furthermore, bone marrow analysis revealed a significant shift towards M1 macrophage polarization in SM@CMC-KP101-treated mice, characterized by a significantly elevated proportion of M1-like macrophages (CD45+F4/80+CD11b+CD86+) in both the SM@CMC-KP101(i.t.) (20.1%) and SM@CMC-KP101(i.v.) (19.4%) groups, compared to the PBS control group (12.1%) (Figure 5g, Supplementary Fig19c). In contrast, the proportion of M2-like macrophages (CD45+F4/80+CD11b+CD206+) remained unchanged (Figure 5h, Supplementary Fig19c). Splenic analysis in CT26-bearing mice revealed that the proportion of CD3+ CD8+ IFN+ cells significantly increased in both the SM@CMC-KP101(i.t.) (16.17%) and SM@CMC-KP101(i.v.) (13.4%) groups (Figure 5i, Supplementary Fig19d). Consistent with this enhanced cytotoxic potential, both intratumoral and intravenous photothermal therapy with SM@CMC-KP101 resulted in a preferential expansion of effector memory CD8+ T cells (CD8+ Tem, CD45+CD3+CD8+CD44+CD62L-) to 37.8% and 38.2% (+179.6% and +182.5% vs control, respectively) (Figure 5j, Supplementary Fig19f). The proportion of central memory CD8+ T cells (CD8+Tcm, CD45+CD3+CD8+CD44+CD62L+) remained stable, suggesting a primary impact on the effector arm of the immune response (Supplementary Fig19f, Supplementary Fig20a). In contrast, the proportion of central memory CD4+ T cells (Figure 5k, CD4+ Tcm) increased to 17.9% and 17.5% (+63.2% and +59.2% vs. control, respectively) (Supplementary Fig19e), while the levels of effector memory CD4+ T cells (CD4+Tem) were unchanged (Supplementary Fig19e, Supplementary Fig20b). Furthermore, both SM@CMC-KP101(i.t.) and SM@CMC-KP101 (i.v.) treatments significantly reduced the proportion of splenic CD4+/CD25+/FoxP3+ Treg cell subpopulations, decreasing from 14.2% in the control group to 8.6% and 8.4%, respectively (Figure 5l, Supplementary Fig21). While ECN, VNP20009, and SM-EV also induced some DC activation and T cell presence, likely due to the inherent immunostimulatory properties of their bacterial components, their lack of prodigiosin loading limited their photothermal capabilities and subsequent therapeutic efficacy, suggesting that SM@CMC-KP101 uniquely combines immune activation with targeted photothermal ablation for enhanced antitumor effects.

Consistent with these immunomodulatory effects, longitudinal tumor volume analysis revealed striking therapeutic efficacy in the SM@CMC-KP101 with 808 nm laser cohort, achieving near-complete tumor regression (15.4 mm³ at day 11 vs 1534 mm³ in controls) accompanied by minor weight fluctuations (Figure 5 m,n; Supplementary Fig22). Analysis of the cured mice prior to rechallenge (Day 14) suggested that the potent broad-spectrum cytotoxicity of prodigiosin in SM@CMC-KP101 likely resulted in the collateral clearance of local lymphocytes alongside tumor eradication. To further validate the establishment of long-term immunological memory and functional anti-tumor immunity, we performed a tumor rechallenge assay (Supplementary Fig23). Mice from the SM@CMC-KP101 (i.t.+NIR and i.v.) groups that achieved complete regression were rechallenged with a high dose of CT26 cells (1 ×106/mice) on Day 14 (Supplementary Fig23a). While all naïve control mice (6/6) developed rapid tumors exceeding 200 mm³, only a minority of the cured mice (1/3 in the i.t. group and 1/4 in the i.v. group) showed tumor recurrence (Figure 5o, p; Supplementary Fig23b). Crucially, the long-term landscape at Day 28 revealed a comprehensive memory expansion, characterized by prominent CD8+ central memory (TCM) upregulation alongside elevated CD8+ TEM and CD4+ memory subsets (Supplementary Fig23c-f). This shift confirms that SM@CMC-KP101 successfully facilitates the evolution from immediate cytotoxicity to durable, broad-spectrum systemic immunological memory.

To address potential risks associated with bacterial therapeutic approaches, we comprehensively assessed the safety profile of SM@CMC-KP101 through multiple methodological approaches. Serum inflammatory markers, including calreticulin (CRT) and procalcitonin (PCT) (Supplementary Fig27), remained within safe ranges, similar with those observed with alternative bacterial vectors. Intriguingly, while transient inflammatory responses were detected in initial blood count analyses, these changes were consistent with expected acute immunological reactions and spontaneously normalized within 15 days (Supplementary Fig28). Histological examinations of major organs revealed no treatment-related pathological alterations (Supplementary Fig29). Furthermore, H&E and Masson's trichrome staining of healed skin after photothermal therapy revealed that, compared to PBS controls, SM@CMC-KP101-treated skin exhibited enhanced collagen deposition, maintenance of a continuous epidermis, and preservation of deeper vessel integrity, indicating a limited and self-resolving superficial effect without discernible damage to normal skin (Supplementary Fig30). Critically, post-treatment survival rates were excellent, and body weights remained stable throughout the experimental period. To further rigorously exclude the risk of sustained systemic infection or cytokine storm often associated with bacterial therapies, we additionally monitored the kinetics of neutrophil levels (CD11b+ Ly6G+) in peripheral blood. Although transient elevation was observed at 24 h post-treatment (consistent with acute immune activation), levels declined significantly by day 14 and returned fully to baseline under normal physiological conditions by day 28 (Figure 5q-s; Supplementary Fig31), with no significant difference compared to the PBS control. This kinetic profile confirms that SM@CMC-KP101 elicits a controlled, transient immune response without persistent bacteremia-driven inflammation, supporting the long-term biosafety of this bacterial platform.

Our study demonstrates that SM@CMC-KP101 represents a breakthrough multimodal therapeutic platform that seamlessly integrates precise near-infrared imaging, targeted photothermal ablation, and immunogenic bacterial clearance. By achieving effective tumor eradication with minimal systemic toxicity and simultaneously reshaping the tumor microenvironment, this approach offers a promising strategy for safe and efficient cancer immunotherapy. The synergistic mechanism of SM@CMC-KP101 not only highlights its potential for clinical translation but also provides an effective strategy for precision oncological interventions.

In vivo Evaluation of SM@CMC-KP101 in the B16F10 Tumor Model
Given that F10 melanoma exhibits higher invasiveness, drug resistance, and immune evasion compared to CT2643, making it considered a more challenging disease to cure in clinical settings. To demonstrate the versatility of our system and the consistency of therapeutic effects across different tumor models, we conducted animal experiments using B16F10 tumor-bearing mice. Notably, B16F10 melanoma is commonly studied in C57BL/6J mice, which have black fur. Due to the light-absorbing properties of this pigmentation, using 808 nm laser irradiation at 4.5W—as previously applied in our CT26 model using BALB/c mice—would inevitably induce non-specific photothermal effects. Therefore, for this model, we opted for a purely intravenous approach without laser irradiation.

To evaluate and compare the therapeutic potential of SM@CMC-KP101 against ECN, VNP20009, SMM, SM, and SM@CMC, we performed an efficacy study employing intravenous (i.v.) administration in the B16F10 melanoma model. Following the safety dose of 1×108 CFU/mouse established in our prior CT26 studies in CT26 tumor-bearing BALB/c mice (Supplementary Fig16), which was confirmed to be both effective and safe. Although most literature reports doses of 1×106 to 1×107 CFU/mouse, lower doses yield limited antitumor effects. Therefore, to enable direct therapeutic comparison, we administered the uniform dosage of 1×108 CFU/mouse. We administered PBS, ECN, VNP20009, SMM, SM, SM@CMC and SM@CMC-KP101 intravenously at this dosage on days 0, 3, 6, and 10 (Figure 6a). with mice being euthanized 14 days post-treatment for subsequent analysis. Analysis of the survival curves (Figure 6b) revealed varying tolerance levels in C57BL/6 mice to the administered bacterial strains at the established dosage of 1×108 CFU/mouse; unfortunately, the ECN-treated group exhibited a 50% mortality rate on the first day post-injection, with complete mortality after the second injection, and the VNP20009 group showed a 25% mortality rate on day one, increasing to 83.3% by day two, also reaching complete mortality after the second administration (Figure 6b). Tumor growth was monitored over the 14-day period, and the SM@CMC-KP101 group exhibited a marked reduction in tumor volume compared to the PBS treated groups (Figure 6c, d; Supplementary Fig32). At the study endpoint, the average tumor size in the SM@CMC-KP101-treated group was 138.9 mm3, compared to 511.92 mm3 in the RSM-treated group and 555.37 mm3 in the SM@CMC-treated group. While the non-attenuated control strain SMM demonstrated a longer survival period, all mice in this group dead by day six. In contrast, the attenuated SM strain, along with the CMC-modified SM@CMC and SM@CMC-KP101 groups, exhibited excellent survival rates, with no observed mortality during the 14-day treatment period, and these groups showed minimal body weight fluctuations (Figure 6e). Notably, separate validation experiments showed that single-component treatments (KP101 alone or CMC alone) exhibited negligible anti-tumor efficacy, with tumor growth kinetics indistinguishable from the control group (Supplementary Fig33). Using these baselines, we calculated the Coefficient of Drug Interaction (CDI) to quantify the synergy. Relative to KP101, SM@CMC-KP101 exhibited strong synergy (CDI = 0.2528; threshold < 0.7), whereas its interaction with CMC was additive (CDI = 1.0046). To further assess the targeting efficiency of SM and its modified versions in comparison to ECN and VNP20009, CFU counts were conducted on homogenized normal and tumor tissues from three mice per group one day following intravenous injection. The results revealed that the attenuated SM strain demonstrated comparable tumor-targeting efficiency to both ECN and VNP20009, with a single-dose administration achieving a bacterial load of 1.97×107 CFU/g in the tumor area, while ECN and VNP yielded counts of 2.78×107 CFU/g and 1.96×107 CFU/g, respectively (Figure 6f). The modified groups, SM@CMC and SM@CMC-KP101, exhibited even higher counts of 2.08×107 CFU/g and 8.98×107 CFU/g, respectively. These findings suggest that the SM strain effectively targets tumors, with SM@CMC-KP101 showing the highest targeting efficiency (Figure 6f). Additionally, the SM strain and its modified derivatives showed a trend toward lower residual rates in normal tissues, particularly the heart and liver, compared to ECN and VNP20009, although these differences were not statistically significant (Figure 6f).

Flow cytometric analysis was performed to evaluate the impact of SM@CMC-KP101 on immune cell populations. After 24 hours, dendritic cells (DCs) were isolated from the tumor-draining lymph nodes, with their maturation markers CD80+ and CD86+ assessed. All experimental groups (PBS, ECN, VNP20009, SMM, SM, SM@CMC, and SM@CMC-KP101) showed a significant increase in the proportion of CD11c+ CD80+ CD86+ dendritic cell (Figure 6g; Supplementary Fig34a), CD69+ CD8+ T cell (Figure 6g; Supplementary Fig34b), and CD69+ CD4+ T cell (Figure 6g; Supplementary Fig34c) subsets in the tumor-draining lymph nodes 24 hours post-treatment, resulting from the exposure to bacterial surface antigens. This indicates that the developed SM@CMC-KP101 treatment system, similar to the established ECN and VNP systems, promoted antigen presentation and T-cell activation. Specifically, the SM@CMC-KP101 group displayed CD11c+CD80+CD86+ dendritic cell, CD69+CD8+, and CD69+CD4+ cell subset proportions of 12.25%, 6.93%, and 8.32%, respectively, which were significantly higher than the PBS treated group proportions of 4.09%, 1.78%, and 2.27%, respectively (Figure 6 g,h,i). Bone marrow analyses at 24 hours revealed a significant rise in CD45+CD11b+F4/80+CD86+ myeloid cells, indicating a shift toward a more proinflammatory M1-like phenotype. Specifically, the percentage of M1-like macrophages (CD11b+F4/80+CD86+) in the SM@CMC-KP101 group increased to 15.85%, compared to 8.15% in the PBS control group (Figure 6j; Supplementary Fig34d). In comparison, the ECN and VNP20009 groups showed M1-like macrophage percentages of 9.13% and 9.74%, respectively, indicating no significant activation (Figure 6j; Supplementary Fig34d). Conversely, the proportions of CD11b+F4/80+CD206+ cells (M2-like macrophages) showed no statistically significant difference among all groups (Figure 6k; Supplementary Fig. 34d). These results suggest that the treatment primarily promotes M1 macrophage polarization in the bone marrow early on, with SM@CMC-KP101 inducing a slightly stronger effect than ECN and VNP20009, while exerting no statistically significant effect on M2 macrophage populations.

After observing the short-term immune activation, we shifted our focus to the long-term effects at 14 days post-treatment. Due to the complete mortality observed in the ECN, VNP, and SMM groups, subsequent analyses at 14 days were limited to the PBS, SM, SM@CMC, and SM@CMC-KP101 groups. The proportion of CD8+ T cells expressing IFNγ (CD3+CD8+IFNγ+) increased significantly in the SM@CMC-KP101 group (13.24%), compared to the PBS control group (4.96%), with the SM@CMC group also showing an increase (10.91%) (Figure 6l; Supplementary Fig34e). Moreover, assessment of memory T cell subsets demonstrated a notable expansion of CD45+CD3+CD8+CD44+CD62L- effector memory T cells (CD8+ Tem) in the SM@CMC-KP101 group (25.73%), compared to the PBS control group (11.37%) (Figure 6m; Supplementary Fig34g). In B16F10 mice receiving intravenous SM@CMC-KP101, CD8+ T effector memory (Tem) cells showed a remarkable activation increase of 126.4% compared to the control group. This was accompanied by increases in central memory (Tcm) CD8+ T cells (56.9%) (Supplementary Fig35a; Supplementary Fig34g), CD4+ Tcm cells (29.5%) (Supplementary Fig35b; Supplementary Fig34f), and CD4+ T Tem cells (100.9%) (Figure 6n; Supplementary Fig34f).

In summary, the SM@CMC-KP101 treatment promotes comprehensive immune responses characterized by the maturation of dendritic cells, robust activation of T cells, and M1 macrophage polarization. Notably, the treatment resulted in a significant increase in the proportion of splenic CD8+ IFNγ+T cell subsets, alongside enhanced populations of effector memory T cells (CD8+ Tem) and central memory T cells (CD4+ Tem). Importantly, hematoxylin and eosin (H&E) staining revealed no obvious tissue damage, indicating the excellent biosafety profile of this approach (Supplementary Fig36). Together, these findings underscore the therapeutic potential of SM@CMC-KP101 in eliciting a sustained anti-tumor immune response and improving long-term tumor control in the B16F10 melanoma model.

Mechanistic Insights into the Antitumor Effects of SM@CMC-KP101
To investigate the mechanisms of SM@CMC-KP101-induced tumor cell killing, we performed proteomic analysis and various cellular and molecular assays in B16F10 tumor cells. SM@CMC-KP101 treatment induces significant changes in protein expression, activates a combination of cell death pathways, and ultimately leading to tumor cell killing through a multi-faceted mechanism. Proteomic analysis revealed that SM@CMC-KP101 treatment resulted in the upregulation of 152 proteins and downregulation of 89 proteins (Figure 7a; Supplementary Fig37-40; Supplementary Data 1). Volcano plot analysis of proteomic data (Figure 7a; Supplementary Data 1) showed the overall changes in protein expression and revealed that MAP kinase-activated protein kinase 5 (Mapkapk5, MK5), a key regulator of the HSP27 phosphorylation involved in cell survival and stress response44, 45, was the most significantly downregulated protein, with a decrease of 8.98-fold. In addition, proteomic analysis identified significant upregulation of ROS synthesis pathways (Maff 5.5-fold upregulated; Mafk 2.1-fold upregulated) and oxidative stress markers (Fos 62.1-fold upregulated; Jun 2.49-fold upregulated), prompting us to validate intracellular ROS levels. Due to the fluorescence interference between the drug and the FITC channel, we used the CellROX Deep Red fluorescent dye to quantify ROS production levels. Results from fluorescence microscopy showed a stronger generation of ROS (Supplementary Fig41). Furthermore, it revealed lysosomal dysfunction markers (Ctsz: 61.5% decrease, Npc2: 60.1% decrease, Ctsh: 69.1% decrease), suggesting disruption of lysosomal function. Kyoto Encyclopedia of Genes and Genomes (KEGG) pathway enrichment analysis (Figure 7b) revealed significant activation of pathways related to autophagy, mitophagy, and necroptosis. Specifically, mitophagy and autophagy-related proteins such as Transcription factor Jun (Jun), Microtubule-associated protein 1 light chain 3 alpha (Map1lc3a) and Microtubule-associated protein 1 light chain 3 beta (Map1lc3b) were significantly upregulated. Additionally, proteins associated with necroptosis, including Interferon gamma receptor 1 (Ifngr1), Tumor necrosis factor receptor superfamily member 1A (Tnfrsf1a), and various components of the CHMP family, were also markedly increased (Figure 7b). Cell viability assay (Figure 7c, Supplementary Fig42) demonstrated that autophagy inhibitor (Chloroquine), apoptosis inhibitor (Z-VAD-FMK), necroptosis inhibitor (Nec-1), and ROS scavenger (Trolox) rescued cells from SM@CMC-KP101-induced cell death both in B16-F10 and CT26 cell lines. Specifically, the necroptosis inhibitor Nec-1 increased the survival rate of F10 cells from 28% to 94%, and CT26 cells from 8% to 59%. Additionally, the apoptosis inhibitor Z-VAD raised the survival rate of F10 cells from 28% to 100% and that of CT26 cells from 8% to 72%, indicating the occurrence of apoptosis and necroptosis (Figure 7c). In addition, western blot analyses showed the effects of SM@CMC-KP101 and PG on proteins involved in mitophagy, autophagy, and necroptosis, which aligned with the proteomic findings (Figure 7d and 7b). Specifically, SM@CMC-KP101 and PG treatment increased the levels of PINK1 and LC3-II/I, while decreasing p62 and CTSB, indicating the induction of mitophagy and autophagic flux. Furthermore, SM@CMC-KP101 and PG treatment increased the levels of phosphorylated mixed lineage kinase domain-like protein (p-MLKL), a marker of necroptosis (Figure 7d). Although caspase-3 and cleaved caspase-3 show a decreasing trend, suggesting that caspase-dependent apoptosis may not be the dominant cell death mechanism, the overall impact of apoptosis should not be excluded (Figure 7d). Consistent with these findings, transmission electron microscopy (TEM) revealed swollen mitochondria and a significant presence of vacuoles within tumor cells, along with evidence of impaired mitophagy after SM@CMC-KP101 treatment (Figure 7e). Collectively, these data indicate that SM@CMC-KP101 treatment induces mitophagy and ROS production, activates stress-related pathways, and ultimately leads to cell death via necroptosis, with a potential contribution from apoptosis.

Given the most significant downregulation of Mapkapk5 observed in our proteomic analysis, and the activation of this stress response pathway by SM@CMC-KP101, we next sought to investigate the specific role of Mapkapk5 in SM@CMC-KP101-induced cell death. We designed three siRNAs targeting different Mapkapk5 gene sites (Supplementary Table 6). qPCR analysis confirmed that transfection with each of the three Mapkapk5 siRNAs resulted in over 80% silencing efficiency of MAPK5 mRNA expression (Supplementary Fig43). CCK8 assay (Figure 7f) showed that knockdown of Mapkapk5 significantly decreased cell viability to 35.89%, 42.44%, and 53.10% of WT, respectively, suggesting that Mapkapk5 expression is critical for cell survival. To further elucidate the mechanism by which Mapkapk5 regulates SM@CMC-KP101-induced cell death, we further assessed the impact of Mapkapk5 knockdown on cell apoptosis. Quantification of Annexin V/PI positive cells (Figure 6g; Supplementary Fig44) revealed that Mapkapk5 knockdown increased the percentage of Annexin V/PI double-positive cells from 2.047% to 2.977%, 4.927%, and 3.083%, respectively, indicating an increase in cell apoptosis. Western blot analyses (Figure 7h) showed that MK5 knockdown did not significantly alter the levels of mitophagy markers (p62, PINK1) or necroptosis markers (p-MLKL, MLKL) (Figure 7h), suggesting that MK5 knockdown is not linked to mitophagy or necroptosis. However, Western blot analysis revealed a surprising result: the levels of caspase-3 and cleaved caspase-3 were downregulated following MK5 knockdown (Figure 7h), contrary to what is typically observed during classical apoptosis46, 47. Intriguingly, MK5 knockdown resulted in the same phenotype as SM@CMC-KP101 and PG treatment, where caspase-3 and cleaved caspase-3 were also downregulated. This suggests that MAPKAPK5 downregulation induces cell death via a caspase-3-independent mechanism.

In summary, SM@CMC-KP101 exerts its cytotoxic effects on B16F10 tumor cells through a multifaceted mechanism (Figure 7i). First, SM@CMC-KP101 may induce mitochondrial dysfunction, leading to the generation of reactive oxygen species (ROS). In response, cells attempt to eliminate damaged mitochondria through mitophagy; however, SM@CMC-KP101 appears to interfere with the fusion and degradation of mitophagosomes in lysosomes, resulting in the accumulation of mitophagosomes. Proteomic analysis revealed a significant downregulation of Mapkapk5, which thereby induces caspase-3-independent apoptosis and mirrors the phenotype observed with SM@CMC-KP101 treatment. Collectively, these results suggest that SM@CMC-KP101 induces tumor cell death via a combination of mitophagy-driven necroptosis and caspase-3-independent apoptosis, with Mapkapk5 downregulation playing a critical role.

Methods
Ethical Statement. All animal experiments were performed in strict accordance with the National Institutes of Health (NIH) guidelines for the care and use of laboratory animals. The animal study protocols were reviewed and approved by the Institutional Animal Care and Use Committee (IACUC) of Nanjing University (approval numbers: IACUC-D2202146).

Mice were housed in a specific pathogen‑free facility under controlled conditions: a 12-hour light/dark cycle (lights on from 8:00 a.m. to 8:00 p.m.), an ambient temperature of 18–22 °C, and 50–60% humidity, with unrestricted access to standard chow and water. The maximum permitted tumor volume was 2000 mm3, and this limit was strictly adhered to throughout all experiments. Mice were humanely euthanized if their tumor volume approached or exceeded this limit, or upon reaching the predetermined experimental endpoint of 14 days.

Materials. Prodigiosin standard was purchased from MedChemExpress (Monmouth Junction, USA). Cell Counting Kit-8 (CCK-8), Annexin V-FITC/PI apoptosis detection kit, and Yopro-1-VS-FITC/PI apoptosis detection kit were obtained from KeyGEN Biotechnology (Nanjing, China). Fluorochrome-conjugated antibodies, including BV480 Rat Anti-Mouse CD45 (#752417), FITC Rat Anti-CD11b(M1/70) (#557396), APC Rat Anti-Mouse CD4(RM4-5) (#553051), Ms CD8a PE 53-6.7 100ug (#553032), Ms CD44 BV421E IM7 50ug (#563970), and PE-Cy7 Rat Anti-Mouse CD62L(MEL-14) (#560516) from BD Biosciences; FIXARBLE VIABILITY DYE EF780 (#65-0865-14), and CD206 (MMR) Monoclonal Antibody (MR6F3), PE (#12-2061-82) from eBioscience; Fc Receptor Blocking Solution, Mouse (#abs9477-200T) from Absin; and FITC anti-mouse FOXP3 (#11-5773-82) and anti-mouse CD25-APC (#17-0251-81) from Invitrogen. Finally, CD3-FITC (#10N0204), CD4-PE (#100512), CD8a-Pacific Blue (#100725), CD11c-APC (#117310), CD80-FITC (#104705), CD86-Pacific Blue (#105022), IFN-γ-APC (#505810) and CD69-FITC (#104505) were acquired from BioLegend (San Diego, USA). The following primary antibodies were used for Western blotting: Pink1 (Abmart, PK05715S), Cathepsin D (Beyotime, AF1645), β-Actin (Beyotime, AF2815), LC3B (Beyotime, AB2023), Hsp27 (Beyotime, AF0183), p-Hsp27(S82) (Cell Signaling Technology, 9709), p38 (Beyotime, AF1111), p-p38 (Cell Signaling Technology, 4511T), caspase3 (Abclonal, A19664), cleaved-caspase3 (Abmart, TA7022), Phospho-MLKL(Ser358) (Cell Signaling Technology, #37333), MLKL (Cell Signaling Technology, #37705), RIPK3 (Abclonal, A5431), and p-RIPK3 (Abclonal, AP1260). All antibodies were used according to manufacturer’s recommendations and listed in Supplementary Table 3, 4. The Chromogenic LAL Endotoxin Assay Kit was purchased from Beyotime Biotechnology (Shanghai, China). KP101 peptide (sequence: FFIVIRDRVFRCG) was custom synthesized by Hefei Peptide Library Co., Ltd (Hefei, China). CellROX™ Deep Red fluorescent probe and TCEP hydrochloride were obtained from Thermo Fisher Scientific (Waltham, USA) and Sigma-Aldrich (St. Louis, USA), respectively.

Development of high-prodigiosin producing Serratia marcescens strains via iterative UV mutagenesis and stepwise screening. The wild-type Serratia marcescens strain NRRL B-1481 was purchased from the American Type Culture Collection (ATCC; Catalog No. 14041). UV mutagenesis and iterative screening52 were performed to obtain high‑prodigiosin (PG)–producing Serratia marcescens mutants. Fresh colonies were inoculated into LB broth (10 g/L tryptone, 5 g/L yeast extract, 10 g/L NaCl) and cultured at 30℃ to late exponential phase (~16 h). Cells were harvested, washed twice with sterile saline, and resuspended to OD600=1.0. To establish the lethal curve and define the working dose, 10‑µL aliquots were spread on sterile Petri dishes (lids open) and irradiated under the germicidal UV lamp of a laminar‑flow hood for 0, 10, 30, 50, 70, or 100 s. Immediately after irradiation, suspensions were serially diluted (103–105) in saline, 100 µL was plated on LB agar, and plates were incubated at 30℃ for 16 h protected from light. Lethality was calculated as 1−N/N0 based on colony‑forming units (N0, non‑irradiated control), and the exposure giving ≥80% lethality (approximately 10 s) was used in all subsequent rounds. For round‑1 mutagenesis, UV‑treated cultures were plated, incubated in the dark, and intensely red colonies were picked for primary screening in 24‑well plates (1 mL fermentation medium per well; 30℃, 200 rpm, 30 h). The fermentation medium contained (per liter) sucrose 20 g, CaCl 3 g, peptone 10 g, and glutamic acid 2 g; the same medium was used for plate‑based screening. After fermentation, 2 mL acidified methanol (adjusted to pH 3.0 with HCl) was added to each culture, samples were sonicated for 20 min, and 200 µL of the extract was transferred to 96‑well plates for spectrophotometric quantification at 535 nm. PG concentrations were calculated from a calibration curve prepared with an authentic prodigiosin standard (MedChemExpress, MCE) and converted to titers (g/L). Top producers from the 24‑well screen were re‑screened in 250‑mL Erlenmeyer flasks containing 50 mL of fermentation medium (30℃, 200 rpm, 30 h) to confirm production and stability; promising isolates were stored as 20% glycerol stocks at −80℃. The best‑producing isolate from round 1 was used as the parental strain for round 2, and the top strain from round 2 served as the parent for round 3, repeating the same workflow (dose confirmation—UV mutagenesis—plate selection—24‑well screening—flask re‑screening—quantification). This stepwise strategy yielded the final high-prodigiosin producing strain SM.

Cell lines. The B16F10 melanoma cells (Cat# IM-M002) w were purchased from IMMOCELL (Xiamen, Fujian, China) and CT26 colon carcinoma (ATCC CRL-2638) authenticated cell lines were purchased from ATCC. RAW264.7 (ATCC TIB-71) were purchased from ATCC. Cells were confirmed mycoplasma free. Cells were cultured in incubators at 37℃ with an atmosphere of humidified 5% CO₂. B16F10 and CT26 cells were grown in DMEM supplemented with 10% (vol/vol) fetal bovine serum (FBS), 1×GlutaMax, 1% (vol/vol) MEM non-essential amino acids solution (Gibco-11140050) and 100 U ml⁻¹ penicillin–streptomycin. No commonly misidentified cell lines were used in this study.

Bacterial Culture and Genetic Manipulation. Serratia marcescens JCS11 (herebefore referred to as SMM) was cultured in LB medium to exponential phase (200 rpm, 30℃).Then, bacteria were washed twice with 0.9% NaCl and resuspended in PBS with serial dilution (10,102,103,104,105 and106) for counting. The bacterial count was determined by the corresponding optical density (OD at 600 nm) measured by plate reader (Infinite 200 PRO, USA). At the same time, the number of bacteria was determined by counting the colony-forming units (CFUs) after culturing the seriabacterial dilution on selective LB agar plates at 37℃ overnight.

The msbB gene encoding lipid A modification enzyme was disrupted in SMM via homologous recombination using the suicide plasmid pDM4 (obtained from our laboratory collection)53. To achieve this, Upstream and downstream homology arms flanking msbB were amplified by high-fidelity PCR (PCR primer sequences are provided in Supplementary Table 5) and cloned into SacI/SalI-digested pDM4 (chloramphenicol-resistant) through seamless assembly. The recombinant plasmid was introduced into E. coli S17-λpir for conjugative transfer. Donor (S17-λpir/pDM4-ΔmsbB) and recipient SMM cultures were mixed (3:1 ratio) and co-incubated on LB agar with 0.22 μm membranes (37℃ for 4 hr, then 28℃ for 16 hr). Transconjugants were selected on LB plates containing 50 μg/mL ampicillin (to suppress donor growth) and 50 μg/mL chloramphenicol. Putative single-crossover mutants were subjected to sucrose counter-selection (10% sucrose in low-salt LB) to eliminate plasmid-retaining strains. Double-crossover mutants were confirmed by PCR amplification of the msbB locus and sequencing using the verification primers ΔmsbB-yz-s (5'-GTTCCGCATCTCCTCCAACTTCAAC-3') and ΔmsbB-yz-a (5'-CAACTGCATCGGTTGAAATACCTCTACC-3'). The Attenuate strain was designated Serratia marcescens JC11 ΔmsbB (SM).

Hemolysis Assay
The hemolytic activity of the bacterial strains was evaluated using Columbia blood agar plates. Briefly, the Columbia Blood Agar Base was purchased from Sangon Biotech (BBI Life Sciences, China; Catalog No. B681083) and prepared according to the manufacturer's instructions, supplemented with 5% defibrinated sheep blood. ECN, SMM, and SM strains were cultured in LB liquid broth until the optical density at 600 nm (OD600) reached 1.0. Subsequently, 5 μL of each bacterial suspension was spotted onto the blood agar plates. A 5 μL drop of Triton X-100 (1% v/v) was spotted as a positive control for hemolysis. The plates were then incubated at 37℃ for 12-24 hours. Following incubation, the plates were visually inspected and photographed to evaluate the formation of hemolytic clearance zones.

Bacterial Culture and Genetic Manipulation. Serratia marcescens JC11 (herebefore referred to as SMM) was cultured in LB medium to exponential phase (200 rpm, 30℃).Then, bacteria were washed twice with 0.9% NaCl and resuspended in PBS with serial dilution (10,102,103,104,105 and106) for counting. The bacterial count was determined by the corresponding optical density (OD at 600 nm) measured by plate reader (Infinite 200 PRO, USA). At the same time, the number of bacteria was determined by counting the colony-forming units (CFUs) after culturing the seriabacterial dilution on selective LB agar plates at 37℃ overnight.

The msbB gene encoding lipid A modification enzyme was disrupted in SMM via homologous recombination using the suicide plasmid pDM4 (obtained from our laboratory collection)53. To achieve this, Upstream and downstream homology arms flanking msbB were amplified by high-fidelity PCR (PCR primer sequences are provided in Supplementary Table 5) and cloned into SacI/SalI-digested pDM4 (chloramphenicol-resistant) through seamless assembly. The recombinant plasmid was introduced into E. coli S17-λpir for conjugative transfer. Donor (S17-λpir/pDM4-ΔmsbB) and recipient SMM cultures were mixed (3:1 ratio) and co-incubated on LB agar with 0.22 μm membranes (37℃ for 4 hr, then 28℃ for 16 hr). Transconjugants were selected on LB plates containing 50 μg/mL ampicillin (to suppress donor growth) and 50 μg/mL chloramphenicol. Putative single-crossover mutants were subjected to sucrose counter-selection (10% sucrose in low-salt LB) to eliminate plasmid-retaining strains. Double-crossover mutants were confirmed by PCR amplification of the msbB locus and sequencing using the verification primers ΔmsbB-yz-s (5'-GTTCCGCATCTCCTCCAACTTCAAC-3') and ΔmsbB-yz-a (5'-CAACTGCATCGGTTGAAATACCTCTACC-3'). The Attenuate strain was designated Serratia marcescens JC11 ΔmsbB (SM). The newly generated materials in this study are available from the corresponding author upon reasonable request.

In-vitro thermokilling assay of SM@CMC-KP101. SM@CMC-KP101 (≈1 × 1011 CFU mL-1 and 1 × 109 CFU mL-1) were exposed to 37℃, 45℃, 50℃, 55℃ or 60℃ for 0 min (control), 3 min or 5 min. Samples were cooled on ice for 1 min immediately after heating. Aliquots were serially diluted, plated in triplicate on LB agar, and incubated at 30℃ for 24 h before CFU enumeration.

Determination of Encapsulation Efficiency. Bacterial encapsulation efficiency was determined by quantifying unencapsulated and total bacterial loads. Briefly, CMC-microencapsulated bacteria were centrifuged (8,000 ×g, 15 min), and the supernatant containing unencapsulated cells was serially diluted, spotted onto agar plates (10 μL/spot, triplicate), and incubated for colony counting (CFU). To measure total bacteria, microcapsules were lysed with 0.5 M EDTA (37℃, 30 min), and the released cells were similarly quantified. Encapsulation efficiency (%) was calculated as:
(1− CFU supernatant / CFU total)×100 (1)

Construction of SM@CMC-KP101. Leveraging chemical modification techniques for targeted peptide functionalization, we coated bacterial surfaces with carboxymethyl cellulose (CMC) to introduce reactive groups for further conjugation. First, bacteria were cultured in LB medium at 37℃ with shaking (250 rpm) until reaching the logarithmic growth phase, collected via centrifugation (3000 × g, 4℃, 5 min), and washed twice with PBS to remove residual medium. The bacterial pellet was resuspended in a 10 g/L CMC solution (e.g., MW 700000) and stirred magnetically at 4℃ for 30 min to ensure uniform coating of CMC on the bacterial surface. After incubation, 2× bacterial volume of 0.2 M CaCl₂ was added to crosslink the CMC layer, stabilizing its attachment to the bacterial surface.

Next, the bacterial pellet was collected by centrifugation and resuspended in PBS (same volume as the pellet). EDC (5 μM) and NHS (5 μM) were added to the suspension and incubated at room temperature for 30 min to activate the carboxyl groups on the CMC layer. KP101 peptide were then added to the activated bacterial suspension, allowing the amide bond formation on the bacterial surface. The reaction was carried out at room temperature for 2 h. Prepared peptide-modified bacteria (e.g., KP101@bacteria) were purified by centrifugation (3220 × g, 4℃, 5 min) and washed three times with PBS. To investigate the impact of surface-bound CMC and KP101 on S. marcescens growth, SM, SM@CMC and SM@CMC-KP101 were serially diluted and plated in triplicate on LB agar, and incubated at 30℃ for 24 h before CFU enumeration.

Photothermal Effect Measurements. The photothermal effects of PG solution at different concentrations (5 mM, 2 mM ,1 mM) and in different solutions were evaluated by using 808 nm laser at 4.5 W/cm2 for 180 s. The aqueous suspensions of SM at different concentrations (1×10⁹, 1×10¹⁰, and 1×10¹¹ CFU/mL) under continuous 808 nm laser irradiation (4.5 W/cm²) for 180 s. Moreover, the photothermal stability of PG and SMM for four on/off irradiation cycles were monitored. The temperature was recorded with an infrared thermal camera.

Number of KP101 on bacterial surface. First, a standard curve between fluorescence intensity and the concentration of FITC labeled KP101 was established using a fluorescence microplate reader (Infinite 200 PRO, USA) with excitation at 488 nm and emission at 520 nm. Then, the fluorescence intensity of FITC labeled KP101 on the S. marcescens surface was measured via Infinite 200 PRO, and the molar concentration was calculated using the regression equation of the standard curve. Next, S. marcescens cell number was determined using the spread plate method. The average number of KP101 molecules per bacterial cell was calculated as:
Navr = (c * v * NA) / N (2)
Navr represents the average number of KP101 on the S. marcescens surface, c represents the molar concentration of KP101, v represents the volume of SM@CMC-KP101 for measurement, NA is Avogadro’s constant 6.02 × 1023 mol−1, and N represents the number of VNP cells for measurement.

Endotoxin Detection. Endotoxin levels were quantified using a chromogenic Limulus amebocyte lysate (LAL) assay kit (C0276S, Beyotime Biotechnology) following the manufacturer’s protocol54. Briefly, samples and endotoxin standards (0.01–1.00 EU/mL, prepared via serial dilution of a 20 EU/mL stock in endotoxin-free water) were mixed with LAL reagent and incubated at 37℃ (9–25 min, depending on expected endotoxin range). Chromogenic substrate was added, followed by a second incubation (6 min), and reactions were terminated with acidic buffer (0.4 M HCl). Absorbance at 545 nm was measured after adding stabilization reagents. A linear standard curve (R² ≥ 0.98) was validated, and sample concentrations were calculated within the curve’s range. All steps utilized endotoxin-free consumables, with negative controls ensuring assay validity.

Detection of Cell Apoptosis. Cell apoptosis was detected using an Annexin V-FITC Apoptosis Detection Kit (KeyGEN Biotech Co., Ltd.) following the manufacturer’s protocol. Briefly, B16F10 cells were seeded onto six-well plates at 2.0×106 cells/well for 24 h. Then, SM@CMC-KP101 was added (final concentration=4.0×107CFU/mL), followed by incubation for 8 h. Moreover, PBS, PG and SM@CMC-KP101 were administered in equivalent proportions. After incubation at 37℃ for 8 h, the cells were collected and stained with the Annexin V-FITC apoptosis detection by fluorescence microscopy (ZEISS LSM880) was performed according to the recommended protocol.

Cell Viability Assay (CCK8). To assess the cytotoxic effect of heat-inactivated bacteria on tumor cells in vitro, CT26 and F10 cells were seeded in 96-well plates at a density of 5×103 cells per well and incubated overnight at 37℃ in a humidified atmosphere containing 5% CO2.
SM@CMC-KP101 were cultured to mid-log phase and then heat-inactivated by incubation at 60℃ for 3 minutes. The concentration of the stock solution of heat-inactivated bacteria was 4×109 CFU/mL. The heat-inactivated bacteria were then diluted in cell culture medium to the indicated working concentrations, with a final concentration of 4×107 CFU/mL based on a 1:100 dilution of the stock.
After overnight incubation of cells, the culture medium was replaced with fresh medium containing various concentrations of heat-inactivated SM bacteria (0, 1×107, 2×107, 3×107, 4×107, 5×107 CFU/mL). After 8 hours of incubation, 10 μL of CCK8 reagent (KeyGEN Biotechnology, China) was added to each well, and the plates were incubated for an additional 2 hours at 37℃. The absorbance was measured at 450 nm using a microplate reader (Bio-Rad, USA). Cell viability was calculated as a percentage relative to the untreated control.

Bacterial Internalization Assay. To quantify bacterial internalization by B16-F10 murine melanoma cells, an amikacin protection assay was performed. B16-F10 cells were seeded in 24-well plates at a density of 5×104 cells per well and incubated overnight at 37℃ in a humidified atmosphere containing 5% CO2. The following day, SMM, SM, SM@CMC, and SM@CMC-KP101 were then washed twice with sterile PBS and resuspended to a final concentration of 1×108CFU/mL in DMEM supplemented with 10% FBS.
Cells were infected with each bacterial strain at a multiplicity of infection (MOI) of 20:1 (bacteria:cell). The plates were centrifuged at 500×g for 5 minutes to synchronize the infection and incubated for 1 hour at 37℃ with 5% CO2. Following incubation, the cell culture medium was removed, and the cells were washed three times with sterile PBS to remove non-adherent bacteria. To kill extracellular bacteria, cells were incubated with DMEM containing 50 µg/mL amikacin for 1 hour at 37℃.
After amikacin treatment, the cells were washed three times with sterile PBS. To release internalized bacteria, cells were lysed with 200 µL of 0.1% Triton X-100 in PBS for 10 minutes at 37℃. The lysate was then serially diluted in PBS, and 10 µL aliquots were plated on LB agar plates. The plates were incubated overnight at 37℃, and the number of colony-forming units (CFU) was enumerated. The percentage of internalized bacteria was calculated as (CFU recovered / CFU input)×100. All conditions were performed in triplicate, and data are presented as mean ± standard deviation.

In vitro macrophage activation. M2 macrophages were polarized from Raw264.7 cells and BMDM as previously described55. Briefly, 1×106 Raw264.7 cells were cultured in a 12-well plate in DMEM medium containing IL-4 (20 ng/mL) for 7~8 days to form M2 macrophages. In parallel, bone marrow-derived macrophages (BMDMs) were generated. Bone marrow cells were flushed from the femurs and tibias of C57BL/6J and cultured in petri dishes in DMEM supplemented with 10% FBS, 1% penicillin/streptomycin, and 20 ng/mL M-CSF for 7 days to differentiate into macrophages. After differentiation, BMDMs were treated with 20 ng/mL IL-4 for 48 hours to induce M2 polarization. The obtained M2 macrophages were washed and incubated with PBS, ECN (final concentration=1×106CFU/mL), SM-EV (final concentration=1×106CFU/mL), SM@CMC-KP101 (final concentration=1×106CFU/mL), and PG (final concentration=12.5 nM) for 8 hours, respectively. After 8 h, the media was aspirated and wells were washed six times with sterile ice-cold PBS. Total RNA from cell samples was isolated according to the manufacturer’s instructions. The relative expression of target mRNAs (iNOS, TNFα, CD206, IL10) was normalized to β-actin as an internal control and calculated using the Log (2-ΔΔCt) method. RT–qPCR primer sequences are provided in Supplementary Table 7.

In vitro BMDC stimulation. Bone marrow-derived dendritic cells (BMDCs) were generated from the bone marrow of 6-8-week-old female C57BL/6J mice using a modified version of previously described methods56. Briefly, murine femurs and tibias were aseptically harvested, and bone marrow cells were flushed into culture medium. After centrifugation to remove debris, cells were resuspended in complete culture medium consisting of RPMI 1640 supplemented with 10% FBS (or serum-free alternative), 1% penicillin/streptomycin, and 20 ng/mL GM-CSF. Cells were cultured at 37℃ in a humidified atmosphere containing 5% CO₂, with a half-medium change and replenishment of GM-CSF on day 3. On days 6-8, non-adherent cells were collected and used as BMDCs. BMDCs were stimulated with microbial strains or other experimental conditions, and their activation was assessed by flow cytometry or other functional assays. The obtained BMDCs were washed and incubated with PBS, ECN (final concentration=1 × 106 CFU/mL), SM-EV (final concentration=1 × 106 CFU/mL), SM@CMC-KP101 (final concentration=1×106 CFU/mL), and PG (final concentration=12.5 nM) for 8 hours, respectively. After 8 h, the media was aspirated and wells were washed six times with sterile ice-cold PBS. Flow cytometric analysis was performed to assess BMDC activation. The following antibodies were used: CD11c-APC (#117310), CD86-PE (105022), and CD80-Pacific Blue (104723). Cells were initially gated on the lymphocyte population based on forward scatter (FSC) and side scatter (SSC) characteristics. Doublets were excluded using FSC-H/FSC-A and SSC-H/SSC-A plots. CD11c+ cells were then gated from the single-cell population, and their activation status was determined by co-expression of CD80+ and CD86+.

Animal Models and In Vivo Experiments. Female BALB/c and C57BL/6J mice (6-8 weeks old) were purchased from Hangzhou Ziyuan Laboratory Animal Technology Co., Ltd. To establish syngeneic tumor models, BALB/c mice and C57BL/6J mice were subcutaneously injected into the left axillary region with 1.0×106 CT26 cells and B16F10 cells, respectively, suspended in 100 μL of PBS. Tumor volume was measured using a digital caliper and calculated as volume=length ×width2 ×π/6. Experiments were conducted when the tumor volume reached 100 mm3. Euthanize the mice when the tumor volume is about to exceed 2000mm3 or has already exceeded 2000mm3, or after 14 days of culture.

To evaluate the in vivo photothermal therapeutic efficacy of ECN, VNP20009, SM-EV,SM@CMC-KP101, and PG in CT26 bearing BALB/c were administered via intratumoral injection at 1.0×1010 CFU/mice. After injection, the tumor underwent 808 nm NIR laser irradiation (4.5 W/cm2 for 3 min), and the corresponding temperature change was recorded using a digital infrared thermal camera. Beside, B16F10 bearing C57BL/6J mice were administered via intravenous injection at 1.0×1010 CFU/mice. The antitumor efficacy was evaluated on the fourteenth day following treatment administration of varying durations (0, 3, 6, or 10 days), necessitated by the substantial tumor volume observed in the control cohort. Throughout the 14-day experimental period, systematic measurements of body weight and tumor dimensions were recorded, alongside continuous monitoring of survival rates among tumor-bearing subjects across different treatment modalities. Upon conclusion of the study, the subjects were humanely euthanized, and tumor specimens were harvested and subsequently preserved in 10% neutral buffered formalin. The preserved tissue samples underwent paraffin embedding, sectioning at 5 μm thickness, and hematoxylin and eosin (H&E) staining for histological examination via digital microscopy.

Evaluation of Synergistic Effects. To evaluate the nature of the interaction between the components, the Coefficient of Drug Interaction (CDI) was calculated based on the tumor volumes measured on day 14 post-treatment. The CDI was determined using the formula:
CDI = EAB / (EA * EB) (3)
where AB represents the ratio of the tumor volume in the combination group to that of the control group, and A and B denote the ratios of the tumor volumes in the respective single-agent groups to the control group57. According to established pharmacological standards, a CDI value less than 1 indicates a synergistic effect, a value equal to 1 indicates an additive effect, and a value greater than 1 implies an antagonistic effect.

DC and T-Cell Stimulation in Tumor-Draining Lymph Node (TDLN) in vivo. The mouse model was established as described earlier, and lymph nodes from the tumor-draining regions were excised 24 hours after intravenous injection and subsequent sacrifice of the mice. Aseptically excise the TDLNs and immediately place them in cold complete RPMI-1640 medium (supplemented with 2% FBS). Maintain the lymph nodes on ice or at 4℃ throughout the procedure. Place the lymph nodes in a 70 µm cell strainer positioned over a 50 mL Falcon tube. Add 2-5 mL of cold complete RPMI-1640 to the strainer and gently disrupt the lymph node tissue by pressing it against the mesh using the plunger of a sterile 5 mL syringe (or a rubber pestle) in circular motions. Wash the cell strainer with an additional 2-3 mL of cold PBS to maximize cell recovery. Collect the filtered cell suspension and centrifuge at 300-400 × g for 5 minutes at 4℃. This cell suspension was treated with CD11c-APC (#117310), CD80-FITC (#104705), CD86-Pacific Blue (#105022) antibodies, followed by flow cytometry analysis to detect dendritic cells (DCs). CD11c+ cells were gated on single cells, with activation status evaluated by CD80 and CD86 co-expression. Additionally, the same cell suspension was processed with CD3-FITC (#100204) or CD69-FITC (#104505), CD4-PE (#100512), CD8a-Pacific Blue (#100725) antibodies to identify T cells. CD69+ cells were gated on single cells, then separated into CD4+ and CD8+ populations58. Representative flow cytometry gating methods are summarized in Supplementary Supplementary Fig19 b,e,f.

Macrophage Phenotyping (24 hours) in 24 h post-treated Bone Marrow. The mouse model was constructed as previously described. At 24 hours post-treatment, the femurs and tibias were aseptically excised, and excess tissue was removed. Using a syringe with a 25G needle, the bone marrow was flushed from the bones with cold complete RPMI-1640 medium (supplemented with 2% FBS). The resulting cell suspension was gently pipetted to break up any clumps and then filtered through a 70 µm cell strainer positioned over a 50 mL Falcon tube to remove debris. The cell strainer was washed with an additional 2-3 mL of cold PBS to maximize cell recovery.After collection, the filtered cell suspension was centrifuged at 300-400 x g for 5 minutes at 4℃. The pellet was resuspended in appropriate buffer for further analysis.
For phenotyping, the cell suspension was stained with the following fluorochrome-conjugated antibodies: BV480 Rat Anti-Mouse CD45 (#752417), FITC Rat Anti-CD11b (M1/70) (#557396), and CD206 (MMR) Monoclonal Antibody (PE, #12-2061-82) from eBioscience. Flow cytometry was performed to determine the percentages of M1 (CD45+CD11b+F4/80+CD86+) and M2 (CD45+CD11b+F4/80+CD206+) macrophages in the bone marrow59, 60. Representative flow cytometry gating methods are summarized in Supplementary Supplementary Fig19c.

Comprehensive Assessment of T Cell Subpopulations in spleen. After the treatment finished, spleens were aseptically excised on day 11 following photothermal therapy in the CT26 tumor model and on day 14 after intravenous injection therapy in the B16F10 model. The excised spleens were immediately placed in cold complete RPMI-1640 medium (supplemented with 2% FBS), maintaining their temperature on ice or at 4℃ throughout the procedure. To prepare splenocyte suspensions, each spleen was placed in a 70 µm cell strainer positioned over a 50 mL tube. 2-5 mL of cold complete RPMI-1640 was added to the strainer, and the spleen tissue was gently disrupted by pressing it against the mesh using the plunger of a sterile 5 mL syringe in circular motions. The cell strainer was washed with an additional 2-3 mL of cold PBS or RPMI to maximize cell recovery. The filtered cell suspension was collected and centrifuged at 300-400 × g for 5 minutes at 4℃.
Splenocytes were stained with specific surface markers for flow cytometry analysis. For (a) splenocyte-derived T cell subsets, live CD45+ cells were gated on CD3+ T cells and then subdivided into CD4+ (stained with APC Rat Anti-Mouse CD4, #553051) and CD8+ populations (stained with Ms CD8a PE, #553032). Memory phenotypes were defined within the CD4+ and CD8+ populations as central memory (TCM: CD44+ CD62L+, using Ms CD44 BV421, #563970, and PE-Cy7 Rat Anti-Mouse CD62L, #560516) and effector memory (TEM: CD44+ CD62L-). For (b) splenocyte-derived Treg cells, live CD45+ cells were first gated on singlets and then on CD4+ T cells. Regulatory T cells (Treg) were identified as CD25+ (stained with anti-mouse CD25-APC, #17-0251-81) and FoxP3+ (stained with FITC anti-mouse FOXP3+, #11-5773-82) cells within the CD4+ population. For (c) splenocyte-derived cytotoxic T cell function, after gating on singlets and CD3+ T cells (stained with CD3-FITC, #100204), CD8+ T cells were identified and further analyzed for intracellular IFN-γ expression using IFN-γ-APC (#505810) following stimulation. Representative flow cytometry gating methods are summarized in Supplementary Supplementary Fig19a, d, g.

Detection of Cellular ROS Levels. ROS levels were determined using a CellROXTM Deep Red Flow Cytometry Assay Kit (ThermoFisher) following the manufacturer’s protocol. The treatment processes were the same as those in the apoptosis assay. Then, the medium was removed, and the cells were washed 3 times with PBS and probed with 5 μM CellROXTM Deep Red in serumfree DMEM. After incubation in 5% CO at 37℃ for 30 min, the cells were collected, and fluorescence was detected by using the PE-Texas Red channel of a flow cytometer (Agilent, NovoCyte).

SiRNA-Mediated Gene Silencing of MAPKAPK5. B16-F10 melanoma cells were cultured in DMEM supplemented with 10% fetal bovine serum (FBS) and penicillin-streptomycin at 37℃ in a humidified atmosphere containing 5% CO2. Three siRNAs targeting MAPKAPK5 were synthesized by GenScript, and the specific sequences are provided in Supplementary Table 6. Transfection was performed using Lipofectamine™ 3000 (Thermo Fisher Scientific) according to the manufacturer's instructions. Cells were seeded in 6-well plates at 1.0 ×105 cells and incubated overnight. For each well, 50 nM of siRNA was mixed with 2 µL of transfection reagent in 200 µL of Opti-MEM and allowed to incubate for 10 minutes at room temperature before being added to the cells. The cells were then incubated for an additional 48 hours at 37℃ with 5% CO2.
To validate the efficiency of MAPKAPK5 gene silencing, total RNA was extracted from the cells using EZ-10 Total RNA Mini-Preps Kit (Sangon Biotech, NO. B618583) following the manufacturer's protocol. Complementary DNA (cDNA) was synthesized using the MightyScript First Strand cDNA Synthesis Master Mix (Sangon Biotech, NO. B639251). Quantitative PCR (qPCR) was conducted using MAPKAPK5 primers purchased from Beyotime, with expression levels normalized to β-actin. The relative expression levels were quantified using the ΔΔCT method.

DIA proteomic analysis
B16F10 cells were seeded onto six-well plates at 2.0 × 10⁶ cells/well for 24 h. Then, PBS or SM@CMC-KP101 was added (final concentration = 4.0×10⁷ CFU/mL; n = 3 independent biological replicates per group), followed by incubation for 12 h. After incubation at 37℃ for 12 h, the cells were collected and frozen for subsequent analysis. Frozen samples (approximately 100 mg) were quickly ground into a fine powder in liquid nitrogen and subsequently lysed in 300 µL lysis buffer supplemented with phosphatase inhibitors and 1 mM PMSF. The samples were further lysed with sonication (1s on/1s off intervals at 80 W for 2 minutes on ice). After sonication, samples were centrifuged at 12,000 × g for 10 min at 4℃ to remove insoluble particles. Protein concentrations were determined using the BCA assay. For SDS-PAGE analysis, an equal amount of protein from each sample was separated by 12% SDS-PAGE gels and stained with Coomassie Brilliant Blue to evaluate sample quality.
Based on the protein concentrations, 50 μg of protein from each sample was diluted to a uniform concentration and volume. Samples were reduced with 5 mM DTT at 55℃ for 30 minutes, cooled to room temperature, and alkylated with 10 mM iodoacetamide in the dark for 15 minutes. Proteins were then precipitated with 6 volumes of pre-cooled acetone at -20℃ overnight and centrifuged at 8,000 × g for 10 minutes at 4℃. The precipitate was redissolved in 50 mM NH₄HCO₃ and digested with Trypsin (1:50 w/w, Trypsin:protein) at 37℃ overnight. Digestion was stopped by adjusting the pH to approximately 3 with phosphoric acid, and samples were desalted using SOLA™ SPE 96-well plates. After drying under vacuum, the samples were spiked with iRT standard peptides (Biognosys) at a 1:20 v/v ratio prior to MS analysis.
Proteomic analyses were performed by Shanghai OE Biotech Co., Ltd. Peptides were separated using a Vanquish Neo UHPLC system (Thermo Fisher Scientific). Mobile phase A consisted of 0.1% formic acid in water, and mobile phase B consisted of 0.1% formic acid in 80% acetonitrile. The separation was performed using a high-throughput gradient strategy (total 22.6 min): 4% B for 0.5 min, 8% B at 0.6 min, 8.5% B at 0.9 min, 22.5% B at 13.9 min, 35% B at 20.8 min, 55% B at 21.2 min, and holding at 99% B from 21.7 to 22.6 min. The flow rate was dynamically adjusted from 1.3 μL/min to 0.8 μL/min during separation and 2.5 μL/min during the washing phase.
The separated peptides were analyzed using an Orbitrap Astral mass spectrometer (Thermo Fisher Scientific) in Data-Independent Acquisition (DIA) mode. The MS parameters were set as follows: Orbitrap resolution at 240,000; full MS scan range of 380-980 m/z; MS/MS scan range of 150-2000 m/z; isolation window of 2 m/z; and higher-energy collisional dissociation (HCD) collision energy at 25%. The cycle time was set to 0.6 s.
For data parsing and protein identification, the raw DIA data were processed using DIA-NN software. The spectra were searched against the UniProt Mus musculus database (release 2024.2.1, combined with iRT sequences). The search parameters were set as follows: trypsin was specified as the cleavage enzyme with a maximum of 1 missed cleavage allowed; carbamidomethylation on cysteine was set as a fixed modification, while oxidation of methionine and acetylation of the protein N-terminus were set as variable modifications. Both the peptide-spectral match (PSM) false discovery rate (FDR) and the protein-level FDR were strictly controlled at ≤ 1% (0.01).

Statistics & Reproducibility
All quantitative data are presented as the mean ± standard deviation (SD) from at least three independent biological replicates unless otherwise specified. No statistical methods were used to predetermine sample sizes; rather, sample sizes for in vivo studies (n≥3) mice per group) were determined based on standards generally accepted in the field and our previous experience with similar tumor models.
For animal experiments, mice were randomly assigned to different treatment groups when their tumor volumes reached approximately 100 mm3. Investigators were blinded to the group allocation during data collection and outcome assessment]. No animals or data points were excluded from the analyses, except for mice that reached the humane endpoint (tumor volume ≥2000 mm3), which were euthanized and recorded as events for survival analysis.
Statistical analyses were performed using GraphPad Prism software (version 10.0). Statistical significance between two independent groups was evaluated using a standard two-tailed unpaired Student's t-test. For comparisons among multiple groups, a two-sided one-way analysis of variance (ANOVA) was used, followed by either Dunnett's post hoc test (for comparing multiple treatment groups to a single control) or Tukey's post hoc test (for multiple pairwise comparisons), as specified in the corresponding figure legends. Animal survival distributions were estimated and plotted using the Kaplan-Meier method. A P value of <0.05 was considered statistically significant. Exact P values are provided in the figure legends.

Peer review rounds
[{"round": 1, "reviewer_comments": {"reviewer_1": "This manuscript describes an engineered Serratia marcescens strain with enhanced prodigiosin production and attenuated virulence, which is applied as a live bacterial vector for anti-tumor immunotherapy and photothermal therapy. The topic is novel and interesting, and the experimental data, while preliminary in some respects, suggest potential value for cancer theranostics.\nHowever, the manuscript contains numerous inconsistencies, ambiguities, and errors—both conceptual and technical—that undermine its scientific rigor and interpretability. In its current form, the manuscript does not meet the standards for publication in a scientific journal. Although the manuscript contains numerous other issues, I have chosen to highlight only a few illustrative examples.\n1. Inconsistent nomenclature and group labeling\nIn lines 271–275, the descriptions of the experimental groups are inaccurate and confusing. For example, SM@CMC-KP101, which refers to the prodigiosin-producing strain, is incorrectly abbreviated again as “SM-PG,” a term previously defined as the non-producing (empty vector) control. Such inconsistencies appear throughout the manuscript and figures, making it difficult for the reader to follow the experimental design.\n2. Memory T cell definition in Figure 4g\nThe memory T cells are labeled as CD3⁺CD8⁺CD44⁺CD62⁺, which is ambiguous. It is unclear whether the authors are referring to central memory or effector memory T cells. Given that these cells are isolated from spleen, the CD62L+ gating in the figure legend is conceptually inconsistent with known tissue localization. Moreover, the figure and its legend are inconsistent in this labeling. The authors must clarify both the phenotype and the anatomical context to support any claims regarding memory T cell activation.\n3. Unsupported illustration in Figure 3k\nFigure 3k illustrates activation of CD8⁺ T cells, yet no corresponding in vitro data are presented in the figure or preceding text to support this claim. While in vivo CD8⁺ T cell activation is shown later (e.g., in Figure 4g), its inclusion in the schematic of in vitro results is misleading unless clearly labeled as hypothetical or supported by data.\n4. Unjustified generalizations\nThe claim that high temperatures “effectively eradicated bacterial carriers” (line 247) is based on in vivo CFU reduction (Figure 4i), but the only in vitro support is qualitative (Figure S4b), showing colony fading and no direct CFU quantification. A more rigorous, quantitative assay would strengthen this conclusion.\n5. Abbreviation use\nNumerous abbreviations appear without being defined on first use (e.g., DA, ECN, VNP20009, PG, etc.), and several are used inconsistently across the manuscript and figures (e.g., SM vs. SMT). A glossary or consistent formatting is needed.\nI recommend that the manuscript be revised thoroughly to address the above concerns. The authors should focus on ensuring consistency in terminology, providing adequate experimental support for mechanistic claims, and improving the clarity and precision of figure legends and group definitions. After these major revisions, the manuscript may be reconsidered for publication.", "reviewer_2": "This study proposes a biohybrid bacterial therapy using engineered Serratia marcescens to achieve high-yield prodigiosin synthesis, tumor-targeted delivery, photothermal ablation, and immunomodulation. The study addresses an innovative topic with a well-designed experimental framework. However, critical data gaps (e.g., dose-response curves, photothermal safety, mechanistic crosstalk) and insufficient discussion of translational challenges must be addressed. The work holds significant potential for publication post-revision.\n1. Clarify the basis for bacterial dosage (e.g., 4×10⁹ CFU) with dose-response experiments (e.g., tumor suppression rates across CFU gradients).\n2. Address potential thermal damage to normal tissues (e.g., skin, vasculature) at 60°C. Include long-term toxicity data or thermal tolerance thresholds.\n3. Supplement Figure 4i with CFU counts in major organs (e.g., liver, spleen) to confirm systemic clearance post-photothermal treatment.\n4. Elucidate the molecular interplay between p38/MK5-HSP27 signaling and mitophagy (e.g., via genetic knockout or pathway-specific inhibitors).\n5. Assess immune memory (e.g., anti-bacterial antibodies) or autoimmune risks post-treatment.\n6. Compare S. marcescens with other bacterial vectors (e.g., VNP20009, EcN) in terms of tumor-targeting efficiency and immune activation to highlight its unique advantages.\n7. Discuss limitations of photothermal therapy (e.g., laser penetration depth, clinical equipment requirements) and challenges in scaling bacterial production.\n8. Benchmark the reported prodigiosin yield (8.3 g/L) against prior studies.\n9. The numbering of the supplementary figures in the supplementary information file appears inconsistent. For example, line 49, Figure S15 is followed by Figure S18, and Figure S18 is followed by Figure S16.\n10. Specify sources of critical reagents (e.g., KP101 peptide synthesis vendor) and include animal ethics approval numbers.", "reviewer_3": "I generally agree with the authors interpretations of the results, and find them of general interest and interest in the area of therapeutic bacteria. They have introduced the use of a bacterium not currently being studied for its anti tumor properties, and also provide a novel image modality with NIR prodigeosin and a novel mode of bacterial elimination. This bacterium was able to kill cells in vitro and had antitumor activity against B16F10 murine melanoma and CT26 murine colon carcinoma models in vivo. This particular melanoma is one that has limited sensitivity to checkpoint inhibitors such as PD-1, which makes that model more difficult to demonstrate a therapeutic effect in with immunomudulators. The bacteria also promoted maturation of dendritic cells, recruitment of T-cells, and favorable macrophage polarization, which may improve the immunotherapeutic effect in these or other models. In B16, apoptosis and mitophagy induced necrosis also occurred and may offer combined advantages in the use of these bacteria.\nHowever there are some specific areas of the conclusions that seem to be implied, but are not specifically addressed.\nThe improved production at a lower temperature in vitro was not demonstrated to occur at a higher temperatures. C57 mice are 36 to 38C and BalbC are 36.4 to 38.4C - how much do they make at mouse temperatures?\nBacterial entry into cells was purportedly shown (see comment for line 155 below), but there was no comparison with the wild type, msbB, or BP101. Is the strain improved in its ability to enter cells, and if so, was that improvement due to the PB101 peptide?\nSimilarly, the attenuated strain with CMC and BP101 was shown to target tumors, but it cannot be determined if that was a preexisting phenotype, or if the msbB, CMC, or PB101 caused any significant improvement.\nRESULTS section\nA discussion of Figure 1 is absent.\nLine 87. The \"mutational breeding\" used in the study is not described.\nLine 89. Physiological temperatures for production of prodigiosin that is the compparitive production of prodigiosin at physiological temperature (37C)? This is especially important if it is down-regulated at higher temperatures (e.g., 42C).\nLine 104 refers to 'Building on the hypoxia-targeting capability of S. marcescens\" while line 116 it states: \"and does not proliferate under hypoxic conditions\". Please explain what appears to be a contradiction.\nThe ability to grow under hypoxic conditions is widely believed to be a component of Clostridium, Salmonella and other bacteria's ability to target tumors, and is not generally linked to toxicity except in the cases of colonizing pre-existing abscesses. How is this an improvement?\nLine 90. How was elevated expression of the PigP operon achieved?\nLine 147-149. I suggest indicating this refers to a schematic diagram as is indicated in Figure Legend 2. Also in Figure 2f, I am unable to read the magnification bars in this version.\nLine 155. Figure 3a needs a higher resolution photo to be convincing as to bacteria having been internalized. Higher magnification and counter staining with a nuclear stain such as DAPI would be helpful. There is also a common assay using gentamicin (assuming your strain is sensitive) DOI: 10.1016/0076-6879(94)36030-8.\nAn additional comment in this regard as stated above: there are no controls for the ability of the wild type to be internalized due to the peptide (e.g, comparison with WT) or the effect of the msbB, CMC coating and BP101 itself on internalization.\nLine 556. Please change tumor models to tumor cells.\nLine 160 and Figure Legend 3c. Which cell line is this?\nFigure Legend 4. Please indicate this is the CT26 tumor model.\nLines 210-211. Please clearly identify the doses being administered by the different routes of injection - e.g., 100 ul of 1x10e11 CFU/ml = 1x10e10 CFU/mouse (or tumor if it).\nLines 221-222. A good tumor/normal tissue ratio is shown, but in the absence of a WT and other comparators, it is unable to be interpreted as an improvement of the wild type. Any such comparison should also individually include the steps in modification, including the delta-msbB, CMC coating, and the addition of KP101.\nFigure Legend 5. Please indicate this is theB16F10 tumor model."}, "author_response": "Response:\nWe thank the reviewer for pointing out the confusion caused by inconsistent abbreviations (e.g., the misuse of \"SM+PG\" for SM@CMC-KP101). In response, we conducted a manuscript-wide audit covering the main text, figure panels/legends, tables, and Supplementary Information. We have replaced all ambiguous or conflicting terms with a single, standardized nomenclature and ensured one-to-one correspondence between each abbreviation and its full experimental definition. To make the study design fully transparent, we now include a summary table (Table S1: Abbreviation-to-sample mapping) listing every abbreviation used in the manuscript and its precise meaning.\nWe believe these revisions eliminate ambiguity and allow the experimental groups to be followed unambiguously throughout the manuscript and figures.\n[...]\nWe thank the reviewer for this insightful comment. We've clarified the specific phenotypes of central memory (Tcm) and effector memory (Tem) CD8+ T cells derived from the spleen in this revised manuscript. Specifically, we redid the in vivo experiments by using mice harboring either CT26 or B16F10 subcutaneous tumors, following treatment via either intratumoral photothermal therapy or intravenous administration. The memory T cells were isolated from the spleens. To provide greater clarity, we have now explicitly defined these T cell populations both upon their initial mention within the manuscript and within the Methods section: Tcm (central memory) is defined as CD3+ CD8+ CD44+ CD62L+, while Tem (effector memory) is defined as CD3+ CD8+ CD44+ CD62L−.\n[...]\nWe appreciate the reviewer's observation that the cartoon in Figure 3k implied CD8+ T-cell activation even though no in-vitro data were provided at that stage of the Results. The intent of this icon was to depict the downstream immune cascade that we later documented in vivo (Figure 4g; Fig 5h,i); however, we agree that its placement inside a figure dedicated to in-vitro findings was misleading.\n[...]\nWe thank the reviewer for highlighting the need for a quantitative in vitro validation. We have now carried out a temperature-controlled study in vitro and in vivo that directly measures colony-forming units (CFU) after short heat pulses that mimic the photothermal conditions generated by our biosynthetic system. We have therefore repeated the heat-eradication experiment under controlled conditions and incorporated rigorous colony-forming-unit (CFU) measurements.\n[...]\nWe appreciate the reviewer for this highly valuable suggestion. We have conducted a manuscript‑wide audit covering the main text, figures, tables, and Supplementary Information, and implemented the following corrections:\nWe have made comprehensive revisions to address these issues, including defining all abbreviations upon first use, ensuring consistent formatting throughout, and adding a glossary for quick reference where appropriate. A detailed summary of these revisions, including specific changes to abbreviations is provided in Table S1.\n[...]\nWe sincerely thank the reviewer for this important question and apologize for the initial lack of clarity and the error in the stated bacterial concentration. In the revision we now provide both (i) in-vitro dose-response data that rationalise the CFU used for cellular assays and (ii) a systematic, 14-day safety/efficacy titration that guided the final intratumoral (i.t.) and intravenous (i.v.) doses used in vivo. All new data are incorporated as Figure 3c (in vitro) and Figure S16 (in vivo) and summarised below.\n[...]\nWe appreciate the reviewer's comments on the potential long-term toxicity. To address this, we have incorporated the following additional data:\n1. Histopathology: H&E and Masson's trichrome staining of major organs and skin tissues post-photothermal treatment.\n2. Bacterial Burden: Quantification of residual bacteria in tissues to assess clearance.\n3. Hematology: Complete Blood Count (CBC) performed 14 days post-treatment.\n4. Systemic Inflammation: Measurement of sepsis-related cytokines (CRT, PCT) to monitor for systemic inflammation.\n[...]\nWe thank the reviewer for these highly valuable suggestions. In this revised manuscript, we performed an in vivo experiment in CT26 tumor-bearing BALB/c mice, quantifying bacterial burdens in major organs following photothermal therapy (PTT). Specifically, mice received an intratumoral injection of 1 × 1010 CFU, followed by 808 nm PTT for 3 minutes. CFU counts in the tumor, heart, lungs, spleen, kidneys, and liver were determined at baseline, day 1, and day 11 post-treatment. These data, demonstrating systemic bacterial clearance, are now presented in Figure 4d, with corresponding raw data available in the Source Data file.\n[...]\nWe thank the reviewer for this insightful suggestion. Following the reviewer's recommendation, we have performed siRNA-mediated MK5 knockdown to solid our mechanistic claims. As suggested, we successfully achieved MK5 knockdown in B16F10 cells, confirmed by qPCR analysis demonstrating a significant reduction in MK5 mRNA levels (Figure S34). Consistent with our proteomics findings, MK5 knockdown resulted in a significant increase in cell death rates (Figure 6f), supporting the idea that MK5 downregulation contributes to the therapeutic effect of our treatment.\n[...]\nWe thank the reviewer for raising this critical point. We have conducted further experiments to assess both immune memory induction and potential autoimmune risks, and the results are now included in the revised manuscript.\n[...]\nWe appreciate the reviewer's highly valuable suggestion. In this revised manuscript, we've expanded our in vivo studies to include a comparative assessment of VNP20009, EcN, SMM, SM, SM@CMC, and SM@CMC-KP101 following intravenous administration in B16F10 tumor-bearing mice (Figure 5).\n[...]\nWe thank the reviewer for raising these critical points regarding the limitations and translational challenges of our approach. This is an essential discussion, and we have now incorporated a detailed discussion into the revised Discussion section of our manuscript. Our response and the corresponding manuscript revisions are structured around the three key areas you identified:\n[...]\nWe appreciate the suggestion to benchmark our prodigiosin titer. As noted in the revised Results and Supplementary Table S2 (summarizing representative production-related studies), the yield of 8.3 g/L in this work is competitive, sitting in the upper range for native S. marcescens under optimized conditions.\n[...]\nWe thank the reviewer for identifying this error. We apologize for the confusion caused by the figure numbering. We have now corrected the numbering for all supplementary figures to ensure they appear in the correct, sequential order. All in-text citations have also been updated accordingly.\n[...]\nWe have added the requested information to the revised Methods section (pp. 21–22) and Supplementary Table 1. A concise summary is supplied below for convenience.\n[...]\nWe thank the reviewer for this insightful comment, as it touches upon a critical aspect of our therapeutic design. As prodigiosin biosynthesis in S. marcescens is temperature-sensitive, our engineered strain is live for pigment production at lower temperatures (28-30 °C), and biosynthesis is negligible at the physiological temperatures found in mice (35-38 °C) (Fig1a, b). To address this, our therapeutic strategy is based on an ex vivo loading model, rather than in situ synthesis.\n[...]\nWe appreciate the request for a head‑to‑head comparison. To determine whether the engineered strain shows improved cellular/tumor entry and whether this is attributable to the KP101 peptide, we conducted matched in vitro studies.\n[...]\nWe thank the reviewer for this insightful comment. To directly address the reviewer's concerns, we performed in vivo experiments to determine the individual contributions of msbB, CMC, and KP101 to tumor targeting. We directly compared SMM, SM, SM@CMC, and SM@CMC-KP101 in tumor-bearing mice.\n[...]\nWe sincerely thank the reviewer for pointing out the absence of a dedicated discussion of Figure 1. We appreciate your meticulous attention to detail.\n[...]\nWe thank the reviewer for highlighting this point and sincerely apologize for the omission of details regarding the \"mutational breeding\" process in the initial submission.\n[...]\nWe thank the reviewer for this question, which addresses the same critical point raised in Q1 regarding prodigiosin production at physiological temperatures.\n[...]\nWe sincerely thank the reviewer for their meticulous reading and for identifying this apparent contradiction. This is a critical point, and we are grateful for the opportunity to clarify the distinction between bacterial tumor targeting and intratumoral proliferation, as these are two separate phenomena key to our strategy.\n[...]\nWe thank the reviewer for this excellent and fundamental question. We completely agree that for classic oncolytic bacteria like Clostridium or certain Salmonella strains, intratumoral proliferation is a primary driver of their therapeutic effect.\n[...]\nWe sincerely apologize for the inaccuracy in our initial submission regarding temperature-controlled prodigiosin (PG) biosynthesis. We are grateful to the reviewer for pointing out this error, which was due to our misinterpretation of the literature.\n[...]\nWe thank the reviewer for these helpful suggestions.We have revised lines 147-149 to indicate that the figure is a schematic diagram, as suggested.\n[...]\nWe sincerely thank the reviewer for this critical and constructive feedback. We agree that the original Figure 3a, a standard bright-field micrograph, was not sufficient to unequivocally demonstrate bacterial internalization versus surface adhesion. To address this, we have performed the experiments suggested by the reviewer and updated the manuscript accordingly.\n[...]\nWe appreciate the reviewer's emphasis on the need for comprehensive controls for internalization. We have addressed this point thoroughly in our response to Q2 and Q3, supported by a new in vitro and in vivo study.\n[...]\nWe appreciate the reviewer’s suggestion. The phrase “tumor models” has been replaced with “tumor cells” to accurately describe the in-vitro experiments.\n[...]\nWe apologize for this omission and appreciate you bringing it to our attention. The experiments shown in Figure 3c were carried out with the murine melanoma cell line B16F10, and we have now clarified this in both the main text and the figure legend.\n[...]\nThank you for noting this. We have updated the Figure 4 legend to clearly indicate that these experiments were performed using the CT26 tumor model.\n[...]\nWe appreciate the reviewer's attention to dosing clarity. We have revised lines 210-211 to provide explicit dosing information:\n[...]\nWe appreciate the reviewer's insightful question regarding the comparison of S. marcescens (SM) with other bacterial vectors. To address this, we compared the survival, tumor growth, and immune response of B16F10 tumor-bearing mice treated with PBS, ECN, VNP20009, SMM, SM, SM@CMC, and SM@CMC-KP101 at the safety dose of 1×108 CFU/mouse (Fig. 5; Figure R23).\n[...]\nThank you for this clarification comment. We have updated the Figure 5 legend to specify that these experiments utilized the B16F10 melanoma tumor model."}, {"round": 2, "reviewer_comments": {"reviewer_2": "The authors have undertaken a comprehensive and rigorous revision of the manuscript. I recommend acceptance of the manuscript in its current form for publication. The revisions have thoroughly addressed the initial concerns, and the work now meets the high standards of the journal.", "reviewer_3": "The authors have substantially improved the manuscript and have satisfactorily answered my original questions.\nIn clarifying the dosing, the need for additional clarification has arisen. The fact that ECN and VNP20009 led to rapid mortality is likely due to their having been used past their tolerated doses (common dose range is 10e6 to 10e7), effectively overdosing the mice with 1x10e8. While the authors may have chosen the dose of 1 x 10e8/mouse as being equivalent in number to the dose of Serratia, the more relevant comparison of drugs for relative efficacy is performed at the effective dose or maximum tolerated dose for each drug. This should be clarified in the text.", "reviewer_4": "I am satisfied with the answers to most of the questions. However, I maintain my criticism regarding the limited scientific rigor and interpretability raised by Reviewer #1. Some of the newly provided data are also unconvincing.\nMemory T cells\nMy impression is that these experiments were performed only once; they should be independently reproduced.\nFigure 3j is described as showing antigen-specific T cells, but no supporting evidence is provided. Tumor-specific T cells should be evaluated as H-2Ld-MuLV gp70 peptide (SPSYVYHQF) tetramer–binding CD8+ T cells (Tet+ CD8+).\nT cells in the draining lymph node were characterized only by general surface markers. Therefore, it is unclear whether they were tumor-specific or potentially bacteria-specific. Why were tumor-infiltrating lymphocytes (TILs) not analyzed?\nFigures 4k–l and 4o–p show an increased memory T-cell population in tumor-eradicated mice, and the authors claim durable antitumor immunity. However, there is no tumor rechallenge study to confirm functional long-term memory T cells generated after bacterial treatment.\nThere is no significant difference in the immune profile between SM@CMC-KP101 (i.t.) and SM@CMC-KP101 (i.v.) (Fig. 4f–m), which is inconsistent with the therapeutic efficacy shown in Fig. 4p (i.t. shows two tumor-eradicated mice, whereas i.v. shows none).\nAdditionally, there are no significant differences in DCs, CD8 T cells, M1 macrophages, M2 macrophages, or Tregs in SM@CMC-KP101 (i.t.)/(i.v.) compared with ECN, VNP20009, and SM-EV. If so, why are there significant differences in antitumor activity? Moreover, neutrophils (CD11b+Gr-1+), which are typically strongly recruited after bacterial infection, were not assessed.\nThe synergy index was not calculated (PMCID: PMC11303714). The combination may be additive rather than synergistic.\nFigure 4b appears to have an incorrect label for SM@CMC-KP101 (DA)."}, "author_response": "Response:\nWe sincerely thank Reviewer #2 for the positive evaluation. Your recognition of the comprehensive, rigorous revisions and support for the manuscript's acceptance in its current form greatly encourages us. We appreciate your affirmation that our work now meets the journal'shighstandards.\n[...]\nThank you for your constructive feedback on our manuscript. We appreciate your insights and would like to clarify our dosing decisions, which have been addressed in the revisedmanuscript:\nWe selected the dose of mouse 1×108 CFU/mouse based on preliminary experiments, whichconfirmedthatthisdoseissafeandessentialforachievingsignificantantitumoreffects with our strain (Serratia). While we recognize that the typical dosing range for ECN and VNP20009is 1×106CFU/mouse to1×107CFU/mouse, our Serratiastrain cantolerate higher doses due to its lower endotoxin levels. This characteristic not only allows for a greater drug load but alsorepresents an important safetysolution for drug deliveryinthe fieldof bacterial therapy,wheredrugloadingcapacityisoftenlimited.\n[...]\nWe appreciate the reviewer’s questionandwouldlike toclarifyour experimentaldesign. The results presented in the initial manuscript were substantiated by multiple independent cohorts rather than a single experiment. Specifically, prior to the comprehensive study, we conducted repeated independent trials to verify the robust therapeutic efficacy of SM@CMC-KP101 (via both intratumoral and intravenous administration) before proceeding to the large-scale, comprehensive study. The final dataset reported included all necessary control groups (ECN, VNP20009, SM-EV) and strictly adhered to randomization protocols, selectingatleastthreebiologicalreplicates(N≥3)pergroupforallimmuneanalysis.\n[...]\nWe thank the reviewer for this critical assessment. We fully agree that characterizing T cells as \"antigen-specific\" requires rigorous validation via tetramer staining (e.g., H-2Ld-MuLV gp70). We acknowledge that our original labeling in Figure 3j was imprecise withoutthisspecificevidence.\n[...]\nWe appreciate the reviewer’s inquiry regarding T cell specificity and TIL analysis. Regardingthe distinctionbetweentumor-specific andbacteria-specificTcells, please refer to our detailed response in Q2, where we discuss the technical limitations and highlight the functional evidence provided by our re-challenge studies. To address the TIL analysis, we performed additional flow cytometry analyses on tumor-infiltrating lymphocytes (TILs) at twokeytimepoints:24hoursandDay14(resultsaddedasFigureS26andFigureS27).\n[...]\nWe acknowledge the necessity of functional validation for the observed memory T cell populations.As detailed in our Response to Q1, we have performed a new tumor rechallenge study to address this point. The results (revised Figure 4 o-p) demonstrate that the SM@CMC-KP101 treated mice successfully rejected the re-inoculated tumor cells, confirmingthatthephenotypicincreaseinmemoryTcellstranslatesintofunctionallong-term protection.\n[...]\nWe thank the reviewer for this careful observation regarding the apparent discrepancy betweenthe similar immune profiles and the different therapeutic outcomes.We attribute this difference to the distinct experimental protocols and the resulting mechanisms of action, whichexertvaryingdegreesofdirecttumorcytotoxicity.\n[...]\nWe acknowledge the reviewer’s observation that there were no significant differences in the recruitment of DCs, CD8+ T cells, M1/M2 macrophages, or Tregs between the SM@CMC-KP101 group and the bacterial control groups (ECN, VNP20009, and SM-EV) (Figure4e-l).Thissimilarityisexpectedbecauseallthesegroupsrelyonbacterialvectorsthat possess intrinsic immunogenicity, thereby triggering a comparable baseline activation of the tumorimmunemicroenvironment.\n[...]\nWe sincerely thank the reviewer for this insightful comment. We fully acknowledge the importance of calculating the synergy index (or Coefficient of Drug Interaction, CDI) to accurately define the nature of the interaction within our drug combination. We agree that distinguishing between additive and synergistic effects provides a more robust evaluation of the therapeutic potential.\n[...]\nWe sincerely thank the reviewer for their meticulous attention to detail.We apologize for this oversight regarding the labeling in Figure 4b. We have corrected the label in the revised manuscripttoaccuratelyrepresenttheSM@CMC-KP101(DA)group."}, {"round": 3, "reviewer_comments": {"reviewer_4": "The authors have carried out a thorough and careful revision of the manuscript. I recommend acceptance for publication in its present form. The revisions comprehensively resolve the original concerns, and the study now satisfies the Nat Comms"}, "author_response": ""}]

Decision letter