Decreased Expression by a Chemically Modified siRNA to PTBP1 Seriously Affects Warburg Effect in Colorectal Cancer Cells

ABSTRACT

Cancer cells preferentially rely on aerobic glycolysis, known as the Warburg effect, to support growth and survival. We previously demonstrated that polypyrimidine tract–binding protein 1 (PTBP1) maintains PKM2 dominance by regulating pyruvate kinase isoform splicing, sustaining the Warburg phenotype. PTBP1 suppression shifts metabolism toward PKM1 dominance, enhancing oxidative phosphorylation, reactive oxygen species (ROS) production, apoptosis, and antitumor immunity. Thirteen chemically modified siR‐PTBP1 derivatives targeting either the coding region or the 3′‐untranslated region (3′‐UTR) of PTBP1 mRNA were synthesized and evaluated in colorectal cancer cell lines. Cytotoxicity, protein expression, oxidative stress, and metabolic alterations were assessed using cell‐based assays, immunoblotting, and metabolomic analysis. siRNA stability was evaluated following nuclease exposure and quantified by TaqMan RT‐qPCR. siRNAs targeting the 3′‐UTR more effectively suppressed PTBP1 expression and increased the PKM1/PKM2 ratio than coding‐region–targeting siRNAs. Among them, derivative 2‐6 showed the strongest cytotoxicity with oxidative stress and apoptosis. Metabolomic profiling demonstrated altered glycolytic flux and preserved pentose phosphate pathway intermediates with activated redox responses, while adenylate and guanylate energy charges remained viable, indicating metabolic stress without energy collapse. Tricarboxylic acid cycle metabolites were elevated, consistent with enhanced oxidative phosphorylation. Derivative 2‐6 showed resistance to nuclease‐mediated degradation. Direct PKM2 knockdown did not induce comparable cytotoxicity. Chemically modified siR‐PTBP1, particularly derivative 2‐6, induces a metabolically vulnerable state characterized by oxidative imbalance, leading to apoptosis. These findings identify PTBP1 as a key regulator of the Warburg effect and support siRNA‐based metabolic targeting as a therapeutic strategy.

Chemically modified siR‐PTBP1 disrupts the Warburg effect by reprogramming pyruvate kinase splicing from PKM2 to PKM1. This metabolic shift activates the TCA cycle, increases ATP production, and excessive ROS generation, and ultimately induces apoptotic cell death in colorectal cancer cells.

Abbreviations

AEC

adenylate energy charge

ATP

adenosine triphosphate

CRC

colorectal cancer

CVS

Crystal Violet Staining

GEC

guanylate energy charge

NGS

next‐generation sequencing

PBS

phosphate‐buffered saline

PC

positive control

PPP

pentose phosphate pathway

ROS

reactive oxygen species

TCA

tricarboxylic acid

1. Introduction

Recent advancements in multimodal cancer therapies have increased the overall 10‐year survival rate of patients with cancer to 59.4% (National Cancer Center, Cancer Information, 2021). However, the prognosis of recurrent and refractory cancers remains poor, highlighting the need for novel therapeutic approaches in addition to existing treatment strategies and drug development.

Recent genome analysis using next‐generation sequencing (NGS) has revealed that genetic mutations in oncogenes vary not only among patients with the same tumor type but also within different tumor regions in a single patient. Consequently, conventional molecular targeted drugs that focus on a single oncogene have limited efficacy and are prone to induce drug resistance. Metabolomic analyses have shown that cancer cells predominantly rely on glycolysis rather than the tricarboxylic acid (TCA) cycle for ATP production, irrespective of oxygen availability (Warburg effect, Figure 1). The Warburg effect has three primary functions: (1) it enhances nucleic acid synthesis via the pentose phosphate pathway, driven by glycolysis‐derived pentoses; (2) it suppresses reactive oxygen species (ROS) production from oxidative phosphorylation in mitochondria, thereby promoting cellular survival; and (3) it modulates tumor immunity by secreting lactate into the extracellular environment, thereby influencing macrophage activity [1, 2].

FIGURE 1.

Proposed model based on previous studies and the present findings of PTBP1‐mediated metabolic reprogramming in colorectal cancer. This schematic illustrates a conceptual model based on our findings and previous studies. In colorectal cancer cells, PTBP1 regulates alternative splicing of pyruvate kinase pre‐mRNA, shifting the balance toward cancer‐associated PKM2 expression. Increased PKM2 expression is associated with enhanced aerobic glycolysis, leading to enhanced glucose uptake, lactate production, biosynthetic precursor generation, and tumor progression. In contrast, PTBP1 downregulation is proposed to promote PKM1 expression and restore mitochondrial oxidative phosphorylation (OXPHOS), thereby partially reversing the Warburg phenotype. Solid arrows indicate experimentally supported pathways, whereas dashed arrows indicate hypothetical mechanisms.

We have previously identified that Polypyrimidine Tract‐Binding Protein 1 (PTBP1) plays a crucial role in regulating the gene expression of Pyruvate Kinase Muscle 1/2 (PKM1 and PKM2), which is essential for maintaining the Warburg effect [3, 4]. PTBP1 functions as a splicing regulator for PKM1 and PKM2, thereby promoting PKM2 expression and sustaining the Warburg effect. As shown in Figure 1, the suppression of PTBP1 expression shifts energy metabolism from PKM2 dominance to PKM1 dominance, resulting in glycolytic inhibition, decreased nucleic acid biosynthesis, and suppressed cancer cell proliferation. This metabolic shift redirects energy production toward the TCA cycle, leading to ROS generation and the subsequent induction of autophagy or apoptosis‐mediated cell death. Additionally, by reducing the dependence on glycolysis and decreasing lactate production, the tumor microenvironment transitions to a state that enhances antitumor immunity.

Although several studies have attempted to suppress PTBP1 expression using miRNAs or conventional siRNAs [3], these approaches have been limited by insufficient stability and transient activity, which restrict their therapeutic applicability [5, 6]. Early PTBP1‐targeting studies primarily employed conventional siRNAs directed against coding regions of PTBP1, although 3′UTR‐targeting siRNAs have also been investigated [3].

Chemical modification of siRNAs has been widely used to improve nuclease resistance, enhance stability in biological environments, and reduce off‐target effects, thereby increasing their potential for therapeutic development [7, 8]. Targeting the 3′‐untranslated region (3′‐UTR) may provide greater sequence specificity, particularly when the target sequence is absent from related genes such as PTBP2 and PTBP3. The target sequence within the 3′‐UTR of our PTBP1‐targeting siRNAs is absent from the corresponding regions of the closely related genes PTBP2 and PTBP3. In addition, the target sequence within the PTBP1 3′‐UTR is identical to the corresponding sequence in mice, indicating that this region is highly conserved between humans and mice.

This study aimed to clarify the role of PTBP1 in regulating the Warburg effect by evaluating an optimized, chemically modified siRNA derivative with enhanced PTBP1‐silencing efficiency.

2. Materials and Methods

2.1. Clinical Samples

Colorectal cancer (CRC) tissues and normal colorectal epithelial cells (isolated from non‐tumorous areas) were obtained from patients who underwent surgery for CRC at the Department of Gastroenterological Surgery, Gifu University Hospital (Gifu, Japan). Among the analyzed cases, three patients (Cases 2, 7, and 10) had received preoperative chemotherapy or chemoradiotherapy prior to surgery. All samples were histopathologically confirmed by hematoxylin and eosin staining. Pathological tumor staging was performed according to the Japanese Classification of Colorectal Carcinomas (2018) (Table 1). All samples were immediately harvested after tissue resection, and protein lysates were prepared using a custom lysis buffer composed of 2× phosphate‐buffered saline (PBS) (25 mL), sodium deoxycholate (0.25 g), NP‐40 (500 μg), and SDS (0.05 g) in a final volume of 300 μL. To this buffer, 15 μL of protease inhibitor cocktail for mammalian cell extracts (Nacalai Tesque, Cat# 25955‐24), 9 μL of Phosphatase Inhibitor Cocktail 2 (Sigma‐Aldrich, Cat# P5726), and 9 μL of Phosphatase Inhibitor Cocktail 3 (Sigma‐Aldrich, Cat# P0044) were freshly added at the time of use. All lysates were stored at −80°C until use.

TABLE 1.

Clinicopathological characteristics of patients with colorectal cancer.

	Sex	Age	Location	Borrmann type	T	N	M	Stage	Tumor size	RAS	BRAF	Preoperative treatment
Case1	M	66	Rb	2	3	0	0	IIa	53	Mutant	Wild	—
Case2	F	46	Rb	2	3	0	0	IIa	45	Mutant	Wild	sRT + CAPOX6
Case3	F	47	S	3	3	0	0	IIa	34	Wild	Wild	—
Case4	F	81	D	2	3	0	0	IIa	25	Wild	Mutant	—
Case5	F	84	T	3	3	0	0	IIa	30	—	—	—
Case6	F	75	RS	0‐Is	2	2a	0	IIIb	18	Wild	Wild	—
Case7	F	68	Rb	2	3	0	0	IIa	40	Mutant	Wild	CAPOX3
Case8	M	75	A	2	4a	0	0	IIb	60	Mutant	Wild	—
Case9	M	81	S	2	3	0	0	IIa	45	Wild	Wild	—
Case10	M	49	Rb	2	2	0	0	I	40	Mutant	Wild	CAPOX + BV4

2.2. Immunohistochemistry

Immunohistochemical staining was performed on formalin‐fixed, paraffin‐embedded tissue sections. After deparaffinization and rehydration, antigen retrieval was carried out using HistoVT One (Nacalai Tesque, Kyoto, Japan). Endogenous peroxidase activity was blocked, followed by incubation with normal horse serum (Vector Laboratories, CA, USA) to reduce nonspecific binding. Sections were then incubated with primary antibodies against PTBP1. Detection was performed using the ImmPRESS HRP polymer detection system (Vector Laboratories), and immunoreactivity was visualized using a DAB substrate. The sections were counterstained with hematoxylin.

2.3. Cell Lines and Culture

The human CRC cell lines DLD‐1 and HT29 were obtained from the Japanese Cancer Research Resources Bank (JCRB, Osaka, Japan) and cultured in RPMI‐1640 (Wako Pure Chemical Industries, Osaka, Japan) supplemented with 8% FBS (Sigma‐Aldrich, St. Louis, MO, USA). Cell cultures were maintained at 37°C in a humidified atmosphere of 5% CO₂. Synthesis of chemically modified siR‐PTBP1 derivatives.

Thirteen chemically modified siRNA derivatives were synthesized, chemically modified, and purified using the FASMAC OLIGO software (Kanagawa, Japan).

2.4. Cell Transfection

Cell transfection was performed using Lipofectamine RNAiMAX Reagent (Invitrogen, Carlsbad, CA, USA) according to the manufacturer's instructions. We created derivatives using chemically modified patented siRNAs (PS, F, etc.; Figure S1). Negative control siRNAs were purchased from HSS (Hokkaido, Japan). The siRNAs were used at a final concentration of 3 nM. For comparison, cells were also transfected with siRNAs targeting PKM2 at the indicated concentrations, and protein expression was analyzed by western blotting.

2.5. Evaluation of Nuclease Resistance and Functional Stability of Derivative 2‐1 and 2‐6

To compare the functional stability of siRNA derivatives following exposure to nuclease‐containing conditions, naked siRNAs (control siRNA, derivative 2‐1, and derivative 2‐6) were incubated in a mixture of 50 μL cell culture medium and 50 μL Lipofectamine RNAiMAX transfection buffer at 37°C for 24 h. After incubation, an additional small volume of Lipofectamine RNAiMAX was added and incubated for 10 min at room temperature, and the entire mixture was applied to DLD‐1 cells at final siRNA concentrations of 2, 4, and 6 nM.

Cells were cultured for 72 h, harvested, and viable cell numbers were determined using a hemocytometer. This assay enables the evaluation of functional siRNA stability after prolonged exposure to extracellular nuclease‐rich conditions.

2.6. Crystal Violet Staining (CVS) Assay

Cell viability was evaluated using CVS assay. Human CRC cells (DLD‐1) were seeded in 96‐well plates at a density of 3000 cells/well and allowed to adhere overnight. The following day, cells were transfected with a chemically modified siR‐PTBP1 derivative. After incubation at 37°C for 72 h, the cells were fixed with 25% glutaraldehyde at room temperature for 15 min. The fixative was removed by rinsing with tap water, and the plates were completely air‐dried. The cells were then stained with a 0.05% crystal violet solution for 15 min. Excess dye was washed off with tap water, and the plates were completely air‐dried. The bound dye was solubilized by adding 100 μL of 0.05% sodium dihydrogen phosphate (NaH₂PO₄) in 50% ethanol to each well. Absorbance was measured at 570 nm using a microplate reader (Sunrise Rainbow) and relative cell viability was calculated based on the absorbance values.

2.7. IC ₅₀ Calculation by Cell Counting

For IC₅₀ determination based on direct cell counting, 5 × 10⁴ DLD‐1 or HT29 cells were seeded per well in 6‐well plates. After 24 h, cells were transfected with siRNA and incubated for 72 h. The supernatants were discarded and the cells were washed with PBS. Trypsin (400 μL) was added and incubated for 7 min at room temperature. The cells were then resuspended by gentle pipetting, and the same volume of culture medium was added to neutralize the trypsin. Cells were counted using a hemocytometer or automated counter, and viability was plotted as a percentage relative to control wells (set as 100%) to determine viable cell number and IC₅₀ values.

2.8. Western Blotting Analysis

DLD‐1 and HT29 cells were seeded at a density of 5 × 10⁵ cells per dish in 10‐cm culture dishes containing 5 mL of complete growth medium and allowed to adhere overnight at 37°C in a humidified incubator with 5% CO₂. For most experiments, cells were collected for western blot analysis 72 h after transfection with siRNAs. For time‐course analysis of apoptotic signaling, cells were harvested at 24, 36, 48, and 72 h after transfection. All cells and clinical samples were lysed for 20 min on ice in lysis buffer containing 1% Protease Inhibitor Cocktail (Sigma‐Aldrich). The protein lysis buffer consisted of 10 mM Tris–HCl (pH 7.4), 1% NP‐40, 0.1% deoxycholic acid, 0.1% sodium dodecyl sulfate (SDS), 150 mM NaCl, and 1 mM EDTA. Proteins were separated on 10% and 12.5% polyacrylamide gels (Wako) by SDS‐PAGE and transferred to PVDF membranes (Perkin Elmer Life Sciences, Boston, MA, USA). The membranes were blocked with 5% nonfat dry milk (Cell Signaling Technology, Danvers, MA, USA) and incubated with the desired primary antibody at 4°C overnight.

Primary antibodies against the following immunogens were used: PTBP1 (Cell Signaling Technology), PKM1 (Cell Signaling Technology), PKM2 (Cell Signaling Technology), LC3B (Cell Signaling Technology), cleaved PARP (Cell Signaling Technology), cleaved caspase‐3 (Cell Signaling Technology), cleaved caspase‐8 (Cell Signaling Technology), cleaved caspase‐9 (Cell Signaling Technology), Bcl‐xL (abcam), RIP (abcam), c‐myc (abcam), β‐actin (Cell Signaling Technology). The protein levels were normalized to β‐actin (Cell Signaling Technology).

2.9. Hoechst Staining

Apoptotic nuclear morphology was assessed by Hoechst 33342 staining. Cells were seeded on coverslips in 24‐well plates and incubated under standard conditions (37°C, 5% CO₂). After treatment, the cells were washed with PBS and fixed with 4% paraformaldehyde for 15 min at room temperature. The cells were then permeabilized with 0.1% Triton X‐100 for 10 min and incubated with Hoechst 33342 solution (5 μg/mL in PBS; Thermo Fisher Scientific, Waltham, MA, USA) for 10 min in the dark at room temperature. After staining, the cells were washed with PBS and mounted using antifade mounting medium. Nuclear morphology was observed under a fluorescence microscope using a DAPI filter. Apoptotic cells were identified by condensed or fragmented nuclei.

2.10. Adenosine Triphosphate (ATP) Assay

Cellular ATP levels were measured using a luminescence‐based ATP assay kit (e.g., CellTiter‐Glo Luminescent Cell Viability Assay, Promega, Madison, WI, USA), according to the manufacturer's instructions. Briefly, cells were seeded into 96‐well plates at a density of 5000 cells per well and incubated under standard conditions (37°C, 5% CO₂) for 48 or 72 h. After treatment, an equal volume of the CellTiter‐Glo reagent was added to each well, and the plate was incubated for 10 min at room temperature to allow cell lysis and luminescence stabilization. Luminescence was measured using a microplate reader. The luminescence intensity was directly proportional to the ATP concentration and the results were normalized to those of the control wells.

2.11. Assessment of Extracellular siRNA Stability by RT‐qPCR and Cell Viability Assay

To evaluate the extracellular stability of siRNA molecules in the culture medium, naked siRNAs (derivative 2‐1 and derivative 2‐6) were incubated in complete medium at 37°C for the indicated time periods. Aliquots of the medium were collected and total RNA was extracted using the miRNeasy Serum/Plasma Kit (QIAGEN). The residual amount of siPTBP1 in each sample was quantified by TaqMan RT‐qPCR with a PTBP1‐specific siRNA detection probe (Thermo Fisher Scientific). Ct values were normalized to the input amount at time 0, and relative siRNA levels were calculated using the ΔΔCt method.

To evaluate whether extracellular exposure affected the cytotoxic activity of siRNAs, naked siRNAs (derivative 2‐1 and derivative 2‐6) were incubated in 100 μL of complete culture medium at nominal concentrations of 200, 400, or 600 nM at 37°C for 24 h. Following incubation, the pre‐incubated siRNA‐containing medium was mixed with Lipofectamine RNAiMAX (Thermo Fisher Scientific) according to the manufacturer's instructions and used for transfection into cultured cells in a final volume of 1 mL of complete culture medium. Cell viability was then assessed by direct cell counting as described for the IC50 determination.

2.12. Assessment of Intracellular ROS Levels by Fluorescence Microscopy

Intracellular ROS levels were evaluated using 2′,7′‐dichlorofluorescin diacetate (DCFDA, Sigma‐Aldrich). The cells were seeded at a density of 7 × 10⁴ cells/well in 6‐well plates. After 24 h of incubation, the cells were transfected with siRNA at concentrations of 0 nM (non‐treated; NT), control siRNA (C), or 0.5, 1, 2, or 3 nM, using Lipofectamine RNAiMAX (Thermo Fisher Scientific) according to the manufacturer's instructions.

Observations were performed 36 h after siRNA transfection because preliminary experiments indicated that substantial cell death occurred at 72 h, which could obscure ROS detection owing to the loss of viable cells. After 36 h of siRNA transfection, the culture medium was replaced with 10 μM DCFDA in serum‐free medium. Cells were incubated in the dark for 20 min at 37°C and washed twice with PBS. For the positive control (PC), cells were then incubated with 110 μM tert‐butyl hydroperoxide (TBHP, Sigma‐Aldrich) in PBS for 20 min at 37°C. All other groups (NT‐, C‐, and siRNA‐treated) were incubated with PBS alone under the same conditions. All procedures were performed under light‐shielded conditions using aluminum foil. The cells were washed twice with PBS, and fluorescence images were acquired using a BZ‐X800 fluorescence microscope (Keyence, Osaka, Japan).

Quantitative analysis of ROS fluorescence was performed using the BZ‐X800 Analyzer software (Keyence). For each sample, three representative fields per well were selected from the average areas across the wells, and the mean values of these three fields were used for subsequent analyses. Total fluorescence intensity in each field was measured and normalized to the cell area in the same field. The fluorescence intensity threshold for a positive signal was manually determined by the investigator based on the observation of the NT and C groups, such that 3%–10% of the area was considered positive. Only areas with fluorescence intensities above this threshold were used for analysis.

2.13. Metabolomic Analysis

Metabolomic profiling was performed using capillary electrophoresis time‐of‐flight mass spectrometry (CE‐TOFMS) and CE triple quadrupole mass spectrometry (CE‐QqQMS). Cells were harvested 36 h after siRNA transfection, which corresponded to the time point at which maximal oxidative stress was observed by fluorescence microscopy. Metabolites involved in glycolysis, the TCA cycle, the pentose phosphate pathway, nucleotide metabolism, and redox regulation were quantified. The adenylate energy charge (AEC) and guanylate energy charge (GEC) were calculated as follows: AEC = (ATP + 0.5 × ADP)/(ATP + ADP + AMP) and GEC = (GTP + 0.5 × GDP)/(GTP + GDP + GMP). Metabolite levels were compared between the control and si‐PTBP1 (2‐6)–treated cells.

2.14. Ethics Statement

This study was approved by the Institutional Review Board of Gifu University Hospital (Approval No. 2025‐161). Written informed consent was obtained from all patients in accordance with the Declaration of Helsinki.

3. Results

3.1. PTBP1 and PKM Isoform Expression in Clinical Colorectal Cancer Samples

First, we analyzed the expression of PTBP1, PKM1, and PKM2 in clinical samples from patients with CRC using both western blotting and immunohistochemical (IHC) analysis (Figure 2) to evaluate the Warburg effect, as shown in Figure 1. Among the analyzed cases, PTBP1 was strongly expressed in five out of 10 samples (Cases 1, 2, 7, 8, and 9), indicating a 50% positivity rate. In Cases 1 and 4 through 9, PKM2 expression in the tumor tissue was consistently higher than that of PKM1 and was elevated compared to that in the adjacent normal mucosa. These findings suggest activation of the Warburg effect in a subset of CRC tumors. Immunohistochemical analysis further demonstrated strong nuclear PTBP1 staining in tumor cells, even in cases showing relatively low PTBP1 expression by western blotting, including Case 3, suggesting that PTBP1 is consistently expressed at the cellular level in tumor tissues. Three patients (Cases 2, 7, and 10) underwent preoperative chemotherapy or radiotherapy; however, no obvious trend in PTBP1 or PKM expression associated with preoperative therapy was observed in this limited cohort. Although high PTBP1 expression appeared to be associated with KRAS‐mutant tumors in several cases in this small cohort, analysis of TCGA datasets revealed no significant difference in PTBP1 expression between RAS‐mutant and wild‐type tumors (Figure S2).

FIGURE 2.

Expression profiles of PTBP1, PKM1, and PKM2 in clinical colorectal cancer tissues. Protein expression levels were examined by western blotting in matched tumor (T) and adjacent non‐tumor (N) tissues from 10 patients with CRC. PTBP1 expression was elevated in five of ten tumor samples, whereas PKM2 predominance over PKM1 was observed in the majority of tumors compared to normal mucosa. In addition, immunohistochemical staining demonstrated strong nuclear PTBP1 expression in tumor cells in all examined cases, including those with relatively low PTBP1 expression by western blotting, whereas adjacent normal mucosa showed weak or negligible staining. These findings support the involvement of PTBP1‐mediated splicing regulation in Warburg‐type metabolic reprogramming observed in colorectal cancer.

3.2. Screening of Chemically Modified siR‐PTBP1 Derivatives Targeting PTBP1 mRNA

To investigate the role of PTBP1 in the Warburg effect, we designed 13 chemically modified siR‐PTBP1 derivatives targeting either the coding region or the 3′‐untranslated region (3′‐UTR) of PTBP1 mRNA to identify the most active suppressors (Figure S1). The 3′‐UTR‐targeting siRNAs were designed based on sequence specificity to minimize off‐target effects on related splicing factor genes such as HNRNP1, PTBP2, and PTBP3. In addition, the targeted 3′‐UTR sequence is conserved across species. Chemical modifications were performed based on various patterns of chemically modified miRNA‐143#12s [9, 10, 11]. Chemical modifications were primarily introduced into the guide strand, particularly within the seed region. Western blotting revealed that the five derivatives (2‐3, 2‐4, 2‐6, 2‐8, and 2‐9) substantially reduced PTBP1 protein levels in DLD‐1 cells. Densitometric analysis using ImageJ software indicated that derivative 2‐6 exerted the strongest knockdown efficiency among the tested siRNAs (Figure 3a). Of note, siRNAs 1‐1 to 1‐4 target the coding region, whereas siRNAs 2‐1 to 2‐9 target the 3′‐UTR of PTBP1.

FIGURE 3.

Screening of silencing efficiency and cytotoxicity of chemically modified siR‐PTBP1 derivatives in DLD‐1 cells. (a) Western blot analysis confirmed PTBP1 silencing and corresponding upregulation of PKM1 in response to selected siRNA derivatives. (b) The crystal violet assay demonstrated a dose‐dependent reduction in cell viability, with derivatives 2‐4 and 2‐6 showing marked cytotoxic activity. (c) Dose–response curves revealed IC₅₀ values of approximately 2.5 nM for derivative 2‐1 and 0.4 nM for derivative 2‐6. (d) Densitometric quantification normalized to β‐actin showed a pronounced increase in the PKM1/PKM2 ratio following treatment with derivative 2‐6.

Cell viability screening assays demonstrated that derivatives 2‐4 and 2‐6 significantly suppressed cell growth. These findings were corroborated by reduced colony formation and increased floating, presumably due to the appearance of apoptotic cells. Derivative 2‐6 showed the strongest cytotoxicity and was selected for further analysis (Figure 3b). The IC₅₀ values were approximately 2.49 nM for derivative 2‐1 and 0.38 nM for derivative 2‐6, with the latter being a chemically modified version of derivative 2‐1, underscoring the critical role of chemical modification in enhancing the potency (Figure 3c).

3.3. PTBP1 Suppression Increases the PKM1/PKM2 Ratio and Induces Cytotoxicity in CRC Cells

Western blotting following treatment with 3 nM derivative 2‐6 demonstrated effective suppression of PTBP1 and a marked increase in PKM1 expression. Quantitative densitometric analysis using ImageJ software, normalized to β‐actin, showed that the PKM1/PKM2 ratio was increased by more than twofold compared with the control (Figure 3d).

To assess whether direct PKM2 suppression could reproduce these effects, we transfected DLD‐1 cells with two independent siRNAs targeting PKM2. Although both siRNAs efficiently reduced PKM2 protein levels, they did not induce LC3B‐II accumulation or PARP cleavage, in clear contrast to derivative 2‐6 (Figure S3a). These results indicated that PKM2 knockdown alone was insufficient to trigger cell death, suggesting that PTBP1 suppression acts through broader metabolic reprogramming rather than simple PKM2 inhibition. Consistently, no apparent morphological features of apoptosis were observed (Figure S3b), whereas only moderate growth inhibition was detected (Figure S3c), suggesting a cytostatic effect.

In addition, to explore potential upstream regulators of PTBP1, the relationship between PTBP1 and c‐Myc expression was examined. As shown in Figure S4, c‐Myc expression was reduced following treatment with derivative 2‐6. Furthermore, c‐Myc knockdown also decreased PTBP1 expression, suggesting that c‐Myc may contribute to the regulation of PTBP1 expression in CRC cells.

3.4. Chemical Modification Enhances Nuclease Resistance and Preserves Functional Activity

To further validate the nuclease resistance of chemically modified siRNAs, we first quantified the residual amount of siPTBP1 in the culture medium using TaqMan‐based RT‐qPCR. As shown in Figure 4a, derivative 2‐6 remained detectable at significantly higher levels than derivative 2‐1 throughout the incubation period (p < 0.01), indicating enhanced stability in the culture medium.

FIGURE 4.

Stability and functional persistence of siRNA derivatives under nuclease‐rich conditions. (a) TaqMan‐based RT‐qPCR quantified remaining siRNA following exposure to culture media for up to 24 h, demonstrating that derivative 2‐6 retained significantly higher levels compared to derivative 2‐1. (b) Functional viability assays showed that derivative 2‐6 maintained cytotoxic activity after 24 h of pre‐incubation, whereas derivative 2‐1 failed to maintain silencing efficacy, indicating improved extracellular stability and functional persistence. Naked siRNAs were pre‐incubated in complete culture medium at 37°C for 24 h at the indicated nominal concentrations before transfection with Lipofectamine RNAiMAX.

Next, to examine whether this increased nuclease resistance translated into retained biological activity, naked siRNAs were pre‐incubated in the culture medium for 24 h prior to transfection (Figure 4b). Derivative 2‐6 maintained strong cytotoxic activity across all tested concentrations, whereas derivative 2‐1 showed no significant reduction in viable cell number compared with the control siRNA‐treated group. Consistent with these findings, derivative 2‐6 significantly reduced viable cell numbers compared to both the control and derivative 2‐1 across all concentrations (p < 0.0001).

Together, these findings indicate that the chemical modifications incorporated into derivative 2‐6 not only enhance nuclease resistance but also preserve functional activity after exposure to extracellular degradation conditions.

3.5. Effects of Derivative 2‐6 in HT29 Cells

In addition to DLD‐1 cells, derivative 2‐6 was tested in HT29 CRC cells, which similarly showed significant PTBP1 suppression, a shift in PKM isoform expression, an increased PKM1/PKM2 ratio, and strong cytotoxicity (Figure 5a). IC₅₀ values in HT29 cells were comparable to those observed in DLD‐1, supporting the robustness of derivative 2‐6's effect across different CRC cell lines (Figure 5b).

FIGURE 5.

Validation of siR‐PTBP1 derivative 2‐6 in an additional colorectal cancer cell line. Western blot analysis of HT29 cells showed a significant decrease in PTBP1 and a corresponding increase in PKM1, similar to the results observed in DLD‐1 cells. IC₅₀ values for derivative 2‐6 in HT29 were comparable to those observed in DLD‐1, suggesting reproducibility and robustness across CRC models.

3.6. siR‐PTBP1 Derivative 2‐6 Induces Metabolic Stress, ROS Generation, and Apoptosis

ATP assay results showed a significant time‐dependent increase in ATP production normalized to cell number: at 3 nM, ATP levels increased from approximately 7.0 × 10³ at 24 h to 1.0 × 10⁴ at 48 h, compared with 5.0 × 10³ and 7.0 × 10³ in the respective controls. These findings suggest a partial metabolic shift toward mitochondrial oxidative phosphorylation under conditions of preserved glycolytic and redox‐supporting pathways, which is consistent with the inhibition of the Warburg effect (Figure 6a).

FIGURE 6.

Metabolic reprogramming, oxidative stress, and apoptosis induced by siR‐PTBP1 derivative 2‐6. (a) ATP assay demonstrating metabolic shift toward mitochondrial oxidative phosphorylation. ATP production was quantified at 24 and 48 h after siRNA transfection. Derivative 2‐6 significantly increased ATP levels in a time‐dependent manner compared to the control groups, consistent with a functional shift from glycolysis‐dominant metabolism to OXPHOS. (b) Representative DCFDA fluorescence images showing intracellular ROS levels under standard and TBHP‐stimulated conditions. (c) Quantitative fluorescence analysis revealed significantly increased ROS levels in siRNA‐treated cells compared with controls, particularly under physiological conditions rather than under oxidative challenge, suggesting restoration of mitochondrial function and a metabolic stress response. (d) Hoechst 33342 staining reveals apoptosis induction by siR‐PTBP1 derivative 2‐6. Representative images show nuclear condensation and fragmentation, characteristic of apoptotic morphology, following treatment. The control group retained the normal round nuclei. (e) Proteins associated with extrinsic apoptotic signaling. Western blot analysis of cleaved PARP, cleaved caspase‐3, and cleaved caspase‐8 following treatment with derivative 2‐6. ACTB was used as a loading control. (f) Proteins associated with intrinsic apoptotic signaling. Western blot analysis of Bcl‐xL, RIP, and cleaved caspase‐9 following treatment with derivative 2‐6. ACTB was used as a loading control.

Fluorescence microscopy with DCFDA staining revealed increased intracellular ROS levels at 36 h after siRNA treatment (Figure 6b,c). Interestingly, under normal conditions (without TBHP stimulation), the difference in fluorescence intensity between the non‐treated/control (NT/C) and siRNA‐treated groups was more pronounced than under TBHP‐stimulated (PC) conditions. The results are presented as the relative fluorescence intensity normalized to that of the control group (C), which was set to 1. Among the siRNA concentrations tested, the 2 nM group exhibited the highest ROS levels. This concentration likely represents the threshold at which oxidative stress is maximized before extensive cell death occurs. In contrast, the 3 nM group showed reduced fluorescence, likely because of extensive cell death at this concentration, which may have affected the measurable ROS signal.

Consistent with the increase in oxidative stress, Hoechst 33342 staining revealed condensed and fragmented nuclei in derivative 2‐6‐treated cells, indicating the induction of apoptosis at 72 h after the transfection (Figure 6d). To further confirm apoptotic cell death at the molecular level, we examined apoptosis‐related signaling pathways following treatment with derivative 2‐6. Cleaved caspase‐3 and cleaved caspase‐8 were induced as early as 36 h after transfection, whereas cleaved PARP became more prominent at later time points (Figure 6e). In contrast, decreased Bcl‐xL expression and detectable cleaved caspase‐9 were observed under selected conditions (Figure 6f). These findings suggest possible involvement of both extrinsic and intrinsic apoptotic signaling pathways. In addition, RIP expression was also decreased following treatment with derivative 2‐6 (Figure 6f). Collectively, these findings indicate that siR‐PTBP1 derivative 2‐6 effectively suppresses PTBP1 expression, induces metabolic reprogramming toward oxidative phosphorylation, increases intracellular ROS levels, and ultimately promotes apoptosis.

3.7. Metabolomic Profiling Reveals Oxidative Metabolic Stress Induced by PTBP1 Suppression

To further characterize the metabolic alterations induced by PTBP1 suppression, metabolomic analysis was performed 36 h after derivative 2‐6 transfection, a time point at which growth suppression was not yet evident, whereas maximal oxidative stress was observed by fluorescence microscopy.

As summarized in Figure 7a, siR‐PTBP1 derivative 2‐6 induced broad metabolic alterations involving glycolysis, the pentose phosphate pathway (PPP), the TCA cycle, and redox‐related metabolism at 36 h after treatment, corresponding to the time point of maximal oxidative stress observed by fluorescence microscopy. Although individual glycolytic and TCA cycle intermediates showed heterogeneous changes, with some metabolites accumulating and others remaining unchanged, the overall metabolic profile indicated a metabolically stressed yet viable state rather than an immediate bioenergetic collapse. Notably, metabolites of the TCA cycle were markedly upregulated, including citrate, cis‐aconitate, isocitrate, and 2‐oxoglutarate. This coordinated accumulation of TCA cycle intermediates, together with increased ATP production and ROS levels, suggests enhanced mitochondrial oxidative phosphorylation following PTBP1 suppression. These results indicated that CRC cells were forced into a metabolically demanding state in which oxidative phosphorylation was upregulated, whereas glycolysis was only partially preserved as a compensatory survival pathway.

FIGURE 7.

Metabolic reprogramming induced by siR‐PTBP1 derivative 2‐6 (a) Schematic overview of metabolic alterations induced by PTBP1 suppression at 36 h after siRNA treatment. All metabolomic analyses were performed at this time point, corresponding to the peak of oxidative stress observed by fluorescence microscopy. Arrows indicate relative directional changes in glycolysis, the pentose phosphate pathway (PPP), the TCA cycle, and redox‐related metabolism. PTBP1 suppression induces oxidative stress while partially preserving glycolysis and PPP, resulting in an intermediate metabolically stressed state rather than immediate energy collapse. (b) Quantitative analysis of key metabolite ratios reflecting metabolic and redox balance. The lactate‐to‐pyruvate ratio, NADPH/NADP⁺ ratio, glutathione levels (GSH, GSSG, and GSH/GSSG), adenylate energy charge (AEC), and guanylate energy charge (GEC) were calculated from metabolomic data. Data are shown as mean ± SD. *p < 0.05, **p < 0.01; N.S., not significant.

A quantitative analysis of the key metabolite ratios is shown in Figure 7b. Treatment with derivative 2‐6 resulted in an increased lactate‐to‐pyruvate ratio, although this difference was not statistically significant, suggesting an altered cytosolic redox balance. Several PPP‐related metabolites, including PRPP, were preserved, indicating the maintenance of nucleotide‐supporting pathways following PTBP1 suppression. In parallel, reduced glutathione (GSH) levels significantly increased, whereas oxidized glutathione (GSSG) showed no significant change, resulting in a largely preserved GSH/GSSG balance. Consistent with enhanced oxidative stress, the NADPH/NADP⁺ ratio showed a decreasing trend, suggesting increased NADPH consumption.

Notably, both the AEC and GEC were significantly decreased but remained within a physiologically viable range in siR‐PTBP1‐treated cells, indicating that global bioenergetic failure had not yet occurred despite pronounced metabolic stress.

Taken together, these results indicate that PTBP1 suppression induces a metabolically stressed but viable intermediate state characterized by oxidative imbalance, partial preservation of glycolytic and PPP‐related pathways, and declining yet non‐collapsing energy homeostasis.

Consistent with the metabolomic findings, western blot analysis demonstrated that PKM2 protein levels remained almost unchanged following siR‐PTBP1 treatment, despite a clear increase in the PKM1/PKM2 ratio. This partial preservation of PKM2 expression may represent an adaptive response aimed at maintaining the glycolytic flux and redox balance through glycolysis and the PPP under metabolic stress.

4. Discussion

In this study, we clearly demonstrated that suppression of PTBP1 using chemically modified siR‐PTBP1 derivatives led to a significant metabolic shift in CRC cells, characterized by an increased PKM1/PKM2 ratio and induction of cell death, effects that were not observed with non‐modified siRNAs.

PTBP1 plays a central role in this process by promoting PKM2 expression through splicing regulation of PKM pre‐mRNA [12, 13]. In the present study, western blot and immunohistochemical analysis further demonstrated that PTBP1 is predominantly expressed and localized to the nucleus of colorectal cancer cells, consistent with its known role as a splicing regulator. Interestingly, Case 3, which showed little or no PTBP1 expression by Western blotting, demonstrated positive nuclear PTBP1 staining by immunohistochemistry. This finding suggests that PTBP1 expression may be underestimated by Western blotting alone. In addition, three cases in the clinical cohort had received preoperative therapy, which may have influenced protein expression profiles. However, no consistent pattern associated with treatment status was observed in this small cohort. Further studies using larger, treatment‐naïve cohorts are warranted to clarify this potential confounding factor. This finding supports the biological relevance of PTBP1‐mediated alternative splicing in maintaining the Warburg effect in clinical CRC tissues. Our findings support the notion that siRNA‐mediated downregulation of PTBP1 shifts isoform expression toward PKM1, thereby altering the energy metabolism of CRC cells toward oxidative phosphorylation. This metabolic reprogramming leads to increased intracellular ATP and ROS levels, suggesting the partial engagement of mitochondrial oxidative phosphorylation under conditions of preserved compensatory metabolism. In addition, we observed a reduction in c‐Myc expression following PTBP1‐targeting siRNA treatment, suggesting a possible link between oncogenic signaling and PTBP1‐mediated metabolic regulation.

Interestingly, despite the robust suppression of PTBP1 and the induction of PKM1, Western blot analysis showed that PKM2 expression did not completely decrease in DLD‐1 and HT29 cells (Figures 3d and 5a). This partial persistence of PKM2 may reflect a biological role beyond glycolysis. This interpretation is consistent with the isoform‐specific dependency reported in tumors, where the requirement for PKM1 versus PKM2 is highly context dependent [14]. Furthermore, PTBP1‐PKM2 interactions have been shown to activate oncogenic STAT3 signaling independent of glycolysis, providing a rationale for residual PKM2 expression even under metabolic reprogramming [15]. PKM2 is known to translocate to the nucleus, where it regulates transcription, cell cycle progression, and tumor‐promoting signaling pathways, which are functions that are not compensated for by PKM1. Therefore, cancer cells may retain a basal level of PKM2 as an adaptive mechanism to preserve proliferative signaling, even when metabolic reprogramming is induced [16, 17]. Maintenance of residual PKM2 expression may be biologically important for cancer cell survival, because PKM2 supports glycolytic intermediates required for the pentose phosphate pathway and nucleotide biosynthesis. This interpretation is consistent with our metabolomic findings showing preservation of PPP‐related metabolites despite marked metabolic stress. Hoechst 33342 staining further confirmed apoptotic nuclear morphology. In addition, induction of cleaved PARP and activation of caspase signaling provided molecular evidence supporting apoptosis following PTBP1 suppression. Furthermore, increased ROS production together with decreased expression of the anti‐apoptotic protein Bcl‐xL and activation of caspase‐9 suggest that PTBP1 suppression induces mitochondrial apoptosis through the intrinsic pathway [18, 19]. Consistent with this mechanism, cleaved caspase‐3 and cleaved PARP were markedly increased following PTBP1‐targeting siRNA treatment, providing molecular evidence of apoptotic cell death. Interestingly, activation of caspase‐8 was also observed, suggesting that the extrinsic apoptotic pathway may additionally contribute to PTBP1‐mediated apoptosis. Because caspase‐8 is a key initiator of death receptor‐mediated apoptosis triggered by ligands such as Fas ligand and TNF‐related apoptosis‐inducing ligand (TRAIL), further studies are warranted to determine whether PTBP1 suppression modulates these upstream death receptor signaling pathways [20, 21]. Collectively, these findings suggest that PTBP1 suppression induces apoptosis through both intrinsic and extrinsic signaling mechanisms, thereby contributing to the anti‐tumor effects of PTBP1‐targeting siRNA.

Importantly, PTBP1 is overexpressed in CRC tissues, which coincides with the significant downregulation of miR‐1 and miR‐133b, even at the adenoma stage [3]. These miRNAs directly target the 3′‐UTR of PTBP1, and their reduced expression contributes to PTBP1 overexpression, thereby promoting the Warburg phenotype. This highlights that the miR‐1/133b–PTBP1 axis is a critical factor in early stage colorectal tumorigenesis and is an attractive therapeutic target.

To further elucidate the metabolic state induced by PTBP1 suppression, we performed metabolomic analysis 36 h after siRNA treatment, corresponding to the time point of maximal oxidative stress observed by fluorescence microscopy. However, at this time point, growth suppression by 2‐6 was not observed relative to the control. This analysis revealed an increased TCA cycle, the preservation of intermediates in the PPP, the activation of redox‐related metabolites, and a significant increase in reduced glutathione levels. Notably, adenylate and guanylate energy charges were largely maintained.

Notably, although cellular ATP levels were increased at later time points (48 h), the adenylate energy charge was significantly reduced at 36 h, indicating that ATP abundance and cellular energy status are not equivalent. While the ATP assay reflects absolute ATP levels, the adenylate energy charge represents the balance among ATP, ADP, and AMP and captures excessive ATP turnover. At 36 h, when oxidative stress was maximal, cells appeared to maintain ATP levels at the expense of increased ADP/AMP accumulation, resulting in a reduced energy charge despite preserved viability. In contrast, increased ATP levels at 48 h likely reflect metabolic adaptation and/or selection of surviving cells with enhanced mitochondrial ATP‐producing capacity. Thus, this discrepancy highlights a metabolically stressed but viable intermediate state rather than energetic recovery.

These findings suggest that PTBP1 suppression drives cancer cells into a metabolically stressed but viable intermediate state characterized by oxidative imbalance rather than immediate energy failure. In particular, the marked elevation of TCA cycle metabolites and increased ATP production strongly implied a shift toward mitochondrial oxidative phosphorylation. Excessive mitochondrial ROS generated under heightened oxidative phosphorylation is known to trigger apoptosis when the antioxidant capacity becomes insufficient [22, 23]. Thus, our data support a model in which PTBP1 suppression induces oxidative stress–driven apoptosis without ATP depletion. The preservation of glycolytic and PPP intermediates together with a sustained energy charge supports the existence of a compensatory metabolic response aimed at maintaining redox homeostasis and nucleotide synthesis under stressful conditions. Although the metabolomic profile suggested enhanced mitochondrial oxidative metabolism, metabolomic analysis alone does not directly measure mitochondrial respiratory function or membrane potential. Therefore, additional functional analyses such as mitochondrial respiration or membrane potential measurements would be required to fully characterize mitochondrial activity. In contrast, PKM2 knockdown alone resulted in growth suppression without apoptotic features, suggesting a cytostatic effect possibly mediated by cell cycle arrest.

In this context, the partial preservation of PKM2 expression observed by western blotting analysis may represent a defensive adaptation, allowing cancer cells to retain their glycolytic flux and NADPH‐generating capacity via the PPP. However, this compensatory mechanism appears to be insufficient for counteracting oxidative stress, ultimately leading to apoptosis and/or autophagy [24]. Collectively, the switch from a PKM2‐dominant to a PKM1‐dominant state following treatment with 2‐6 resulted in dysregulated TCA cycle activity and increased oxidative stress, which is a critical event preceding cell death. Consistent with this interpretation, direct PKM2 knockdown failed to induce apoptosis despite a substantial reduction in PKM2 protein, whereas 2‐6 robustly activated LC3B‐II and PARP cleavage (Figure S3a). This finding supports the concept that PTBP1 suppression–mediated isoform switching and network‐level metabolic remodeling, rather than PKM2 inhibition alone, underlies the cytotoxicity observed with 2‐6. To validate this mechanism, we compared the cytotoxic activity of 2‐6 with that of siRNAs targeting PKM2. As expected, 2‐6 exhibited a markedly higher cytotoxic activity than PKM2 knockdown alone, supporting the significance of PTBP1 suppression–mediated PKM isoform switching in dysregulating the Warburg effect.

Among the 13 siR‐PTBP1 derivatives tested, derivative 2‐6 emerged as the most potent, showing the greatest suppression of PTBP1, the highest PKM1/PKM2 ratio, and the strongest cytotoxic effect. These effects were consistent in both DLD‐1 and HT29 CRC cell lines, confirming the broad utility of derivative 2‐6. Furthermore, the metabolic shift away from glycolysis results in altered lactate metabolism, which may improve the tumor immune microenvironment by reducing immune suppression [2].

Importantly, our data showed that siRNAs targeting the 3′‐untranslated region (3′‐UTR) of PTBP1 mRNA were more effective in reducing PTBP1 levels and shifting PKM isoform expression than those targeting the coding region. The 3′‐UTR‐targeted siRNAs may offer enhanced specificity and reduced off‐target effects due to the higher sequence variability and target uniqueness in non‐coding regions, in contrast to the more conserved nature of coding sequences. This strategy highlights a key advantage in designing nucleic acid drugs that exploit the untranslated regions of oncogenic mRNAs for selective inhibition [25, 26].

In addition to sequence targeting, we employed chemical modifications such as 2′‐fluoro (2′‐F) ribose substitution and phosphorothioate (PS) backbone linkage to improve the stability, cellular uptake, and resistance to nuclease degradation of the siRNA molecules. The 2′‐F modification enhances binding affinity to the RNA target while increasing resistance to endonucleases, and the PS modification contributes to both improved serum stability and prolonged half‐life by reducing degradation by exonucleases [27, 28]. These modifications are commonly used in clinical‐grade oligonucleotide therapeutics, and are critical for the in vivo applicability of nucleic acid drugs [29]. A few PS modifications at the ends of the antisense strand may enhance activity.

Furthermore, our nuclease‐resistance assay demonstrated that derivative 2‐6 retained strong cytotoxic activity even after 24 h of pre‐incubation in culture medium, whereas derivative 2‐1 exhibited a loss of activity. These findings provide direct functional evidence that the chemical modifications applied to derivative 2‐6 substantially enhance resistance to extracellular nuclease degradation and allowed siRNA to maintain its biological activity under destabilizing conditions. This enhanced stability is consistent with the superior PTBP1 suppression and cytotoxic potency observed in the main assays and highlights derivative 2‐6 as a more promising candidate for in vivo applications where nuclease exposure is unavoidable. In addition to the functional assay, TaqMan RT‐qPCR analysis demonstrated that derivative 2‐6 remained quantitatively detectable in the culture medium for longer than derivative 2‐1, further supporting its enhanced molecular stability. This molecular‐level evidence is consistent with the functional nuclease‐resistance assay and underscores the advantages of the chemical modifications incorporated into derivative 2‐6.

Fluorescence microscopy analysis using DCFDA staining revealed that siRNA‐induced ROS accumulation was more pronounced under normal conditions (without TBHP stimulation) than under TBHP‐stimulated conditions. Among the siRNA concentrations tested, 2 nM resulted in the highest ROS levels, whereas 3 nM siRNA reduced ROS signaling. We speculate that this decrease was due to extensive cell death at higher concentrations, which reduced the number of viable cells available for ROS quantification. These findings highlight a potential threshold at which oxidative stress leads to irreversible cellular damage, and suggest that moderate siRNA doses may optimally balance metabolic reprogramming and cell viability. Similar dose‐dependent effects of ROS on ROS‐induced apoptosis have been reported in other cancer models [23, 24]. Our findings suggest that combining 3′‐UTR specificity with rational chemical modifications results in an optimized siRNA therapeutic platform capable of selectively modulating cancer‐specific splicing factors such as PTBP1. Preliminary experiments showed that derivative 2‐6 exhibited significant growth suppression in gastrointestinal cancer cell lines, including gastric and esophageal cancer cell lines.

In conclusion, this study demonstrates that chemically modified siRNAs targeting the 3′‐UTR of PTBP1 effectively suppress the Warburg effect in CRC cells by shifting metabolic profiles toward oxidative phosphorylation, thereby highlighting the therapeutic potential of chemical modification. These findings indicate that PTBP1 plays a central role in regulating the Warburg effect. This metabolic reprogramming results in increased ATP production, induction of apoptosis, and decreased cell viability. Derivative 2‐6, in particular, emerged as a potent candidate with high specificity and cytotoxicity in both DLD‐1 and HT29 cells. These findings support the further development of siRNA‐based nucleic acid therapeutics targeting PTBP1 for the treatment of CRC, with potential applicability across various tumor contexts.

Furthermore, the enhanced nuclease resistance observed in derivative 2‐6 supports its potential applicability in in vivo settings, where extracellular degradation is a major limiting factor for siRNA therapeutics.

The proposed mechanism by which chemically modified PTBP1‐targeting siRNA suppresses colorectal cancer cell growth through metabolic reprogramming and apoptosis is summarized in Figure 8.

FIGURE 8.

Graphical abstract. Proposed mechanism of chemically modified siPTBP1 derivative 2‐6. PTBP1 suppression increases the PKM1/PKM2 ratio, inhibits the Warburg effect, enhances oxidative phosphorylation, increases ROS and ATP production, and induces apoptosis in colorectal cancer cells.

Author Contributions

Keita Matsumoto: conceptualization, investigation, formal analysis, writing – original draft, writing – review and editing. Hirokatsu Hayashi: conceptualization, investigation, formal analysis, writing – review and editing. Yoshihisa Tokumaru: formal analysis, writing – review and editing. Seito Fujibayashi: resources. Noriki Mitsui: resources. Masahide Endo: resources. Takeshi Horaguchi: resources. Yuji Hatanaka: resources. Ryoma Yokoi: resources. Chika Mizutani: resources. Masashi Kuno: resources. Masahiro Fukada: resources. Ryuichi Asai: resources. Itaru Yasufuku: resources. Yuta Sato: resources. Jesse Yu Tajima: resources. Yoshihiro Tanaka: resources. Remi Nakashima: investigation, formal analysis. Yukihiro Akao: conceptualization, investigation. Nobuhisa Matsuhashi: supervision, writing – review and editing.

Funding

This work was supported by research funding from Chugai Pharmaceutical Co. Ltd. and eNA‐biotec.

Ethics Statement

This study was approved by the Institutional Review Board of Gifu University Hospital (Approval No. 2025‐161).

Consent

Written informed consent was obtained from all patients.

Conflicts of Interest

The authors declare that this study received research funding from Chugai Pharmaceutical Co. Ltd. and eNA‐biotec. The funder had no role in study design, data collection, analysis, interpretation, or manuscript preparation.

Supporting information

Figure S1: Design strategy and chemical structures of siR‐PTBP1 derivatives targeting coding and 3′‐UTR regions. Thirteen siRNA derivatives were designed based on two target regions of PTBP1 mRNA (coding sequence and 3′‐UTR). Each derivative incorporated phosphorothioate (PS) linkages and/or sugar modifications, including 2′‐fluoro and 2′‐O‐methyl substitutions, to enhance nuclease resistance and silencing potency. Sequences, modification patterns, and classifications (1‐series: coding region; 2‐series: 3′‐region) are shown.

Figure S2: PTBP1 expression according to KRAS/NRAS mutation status in TCGA colorectal cancer cohort. PTBP1 expression levels (Z‐score) were compared between tumors with KRAS/NRAS mutations (Mutant) and those without mutations (Wild) using data from The Cancer Genome Atlas (TCGA) colorectal cancer cohort. No significant difference in PTBP1 expression was observed between the two groups (p = 0.8), indicating that PTBP1 expression is independent of RAS mutation status.

Figure S3a: Effects of PKM2 knockdown on PTBP1‐related signaling and apoptosis. DLD‐1 cells were transfected with siRNA targeting PKM2. Western blot analysis confirmed efficient suppression of PKM2 expression, while PTBP1 expression remained unchanged. No PARP cleavage was detected, indicating that PKM2 knockdown alone was insufficient to induce apoptosis. β‐actin (ACTB) was used as a loading control.

Figure S3b: Morphological changes following PKM2 knockdown. Representative phase‐contrast images of DLD‐1 cells transfected with siPKM2 (3 nM). No apparent morphological features of apoptosis, such as cell shrinkage or detachment, were observed compared with the non‐treated (NT) group.

Figure S3c: Effects of PKM2 knockdown on cell proliferation. DLD‐1 cells were transfected with siRNA targeting PKM2 at the indicated concentrations. PKM2 knockdown resulted in a dose‐dependent reduction in cell proliferation compared with control groups. However, the degree of growth inhibition was moderate, indicating a cytostatic effect rather than induction of cell death.

Figure S4: Effects of PTBP1 suppression on c‐Myc expression. DLD‐1 cells were transfected with siR‐PTBP1 (derivative 2‐6). Western blot analysis showed effective suppression of PTBP1, accompanied by increased PKM1 expression without a marked reduction in PKM2 levels. c‐Myc expression was also decreased following PTBP1 suppression, suggesting that PTBP1 suppression is associated with altered c‐Myc expression. β actin (ACTB) was used as a loading control.

Data Availability Statement

The data that support the findings of this study are available from the corresponding author upon reasonable request.