Unexpected Photocatalytic Degeneration of NAD + for Inducing Apoptosis of Hypoxia Cancer Cells

Abstract

Open AccessCCS ChemistryRESEARCH ARTICLES22 Dec 2022Unexpected Photocatalytic Degeneration of NAD+ for Inducing Apoptosis of Hypoxia Cancer Cells Shengpeng Xia, Haotian Bai, Endong Zhang, Wen Yu, Zhiqiang Gao, Fengting Lv, Yiming Huang, Daoben Zhu and Shu Wang Shengpeng Xia Beijing National Laboratory for Molecular Sciences, Key Laboratory of Organic Solids, Institute of Chemistry, Chinese Academy of Sciences, Beijing 100190 College of Chemistry, University of Chinese Academy of Sciences, Beijing 100049 Google Scholar More articles by this author , Haotian Bai *Corresponding authors: E-mail Address: [email protected] E-mail Address: [email protected] E-mail Address: [email protected] Beijing National Laboratory for Molecular Sciences, Key Laboratory of Organic Solids, Institute of Chemistry, Chinese Academy of Sciences, Beijing 100190 Google Scholar More articles by this author , Endong Zhang Beijing National Laboratory for Molecular Sciences, Key Laboratory of Organic Solids, Institute of Chemistry, Chinese Academy of Sciences, Beijing 100190 Google Scholar More articles by this author , Wen Yu Beijing National Laboratory for Molecular Sciences, Key Laboratory of Organic Solids, Institute of Chemistry, Chinese Academy of Sciences, Beijing 100190 College of Chemistry, University of Chinese Academy of Sciences, Beijing 100049 Google Scholar More articles by this author , Zhiqiang Gao Beijing National Laboratory for Molecular Sciences, Key Laboratory of Organic Solids, Institute of Chemistry, Chinese Academy of Sciences, Beijing 100190 College of Chemistry, University of Chinese Academy of Sciences, Beijing 100049 Google Scholar More articles by this author , Fengting Lv Beijing National Laboratory for Molecular Sciences, Key Laboratory of Organic Solids, Institute of Chemistry, Chinese Academy of Sciences, Beijing 100190 Google Scholar More articles by this author , Yiming Huang *Corresponding authors: E-mail Address: ba[email protected] E-mail Address: [email protected] E-mail Address: [email protected] Beijing National Laboratory for Molecular Sciences, Key Laboratory of Organic Solids, Institute of Chemistry, Chinese Academy of Sciences, Beijing 100190 Google Scholar More articles by this author , Daoben Zhu Beijing National Laboratory for Molecular Sciences, Key Laboratory of Organic Solids, Institute of Chemistry, Chinese Academy of Sciences, Beijing 100190 Google Scholar More articles by this author and Shu Wang *Corresponding authors: E-mail Address: [email protected] E-mail Address: [email protected] E-mail Address: [email protected] Beijing National Laboratory for Molecular Sciences, Key Laboratory of Organic Solids, Institute of Chemistry, Chinese Academy of Sciences, Beijing 100190 College of Chemistry, University of Chinese Academy of Sciences, Beijing 100049 Google Scholar More articles by this author https://doi.org/10.31635/ccschem.022.202202463 SectionsSupplemental MaterialAboutAbstractPDF ToolsAdd to favoritesDownload CitationsTrack Citations ShareFacebookTwitterLinked InEmail Developing customized chemical reactions that could regulate a specific biological process on demand is regarded as an advanced and promising strategy for treating diseases. However, conventional chemical reactions become challenging in complex physiological environments, which demand mild reaction conditions, high efficiency, good biocompatibility, and strong controllability. Moreover, the effects of the achieved reactions on the real biological system are usually further lessened. Herein, we describe an advanced photocatalytic reaction that irreversibly converted nicotinamide adenine dinucleotide (NAD+) to nicotinamide and adenosine diphosphate (ADP)-ribose by the cationic conjugated poly(fluorene-co-phenylene) (PFP). This reaction was introduced to tumor cells and triggered cell apoptosis. Under white-light illumination, the photocatalytic reaction decreased the NAD+ ratio in tumor cells, disrupted the mitochondrial membrane potential, inhibited the synthesis of adenosine triphosphate (ATP), and effectively induced apoptosis. We propose a mechanism of the reaction where PFP is photoexcited to PFP*, and the obtained photoelectrons are transferred from PFP* to NAD+ to produce nicotinamide and another unstable intermediate, ADP-ribosyl radical. ADP-ribosyl radical quickly reacts with triethanolamine to form ADP-ribose. This intracellular utilization of a specific photocatalytic reaction could offer a new approach to affect biological function for efficient cancer treatment. Download figure Download PowerPoint Introduction Life is based on continuous cellular metabolism processes, where many substrates and products are often involved in various chemical reactions.1,2 These synergetic reactions, including intracellular redox, enzyme cascade, and reversible reaction, maintain homeostasis of the biological system in individual bodies.3 However, biological metabolic processes with erroneous chemical reactions can cause metabolic disorders and dysfunction, even injury, cancer, and autoimmune disease.4–7 Customized chemical reactions that could regulate specific biological processes are regarded as an advanced and promising strategy for treating diseases.8 However, the complex physiological environments, including the near-neutral pH, mild temperature, pressure, and hypoxic microenvironment, also put many preconditions on developing these reactions.9 The additional demands of excellent controllability and biocompatibility make these investigations more challenging.10 Moreover, cellular self-healing ability11 would further weaken the effects of these reactions because living cells prefer to maintain a chemical balance in the body. Cancer, with high morbidity and mortality, has significantly threatened human life.12–14 Hypoxic microenvironments often cause worse disease outcomes by protecting cancer cells against apoptosis and enhancing the ability of tumor metastasis.3,15,16 Nicotinamide adenine dinucleotide (NAD) is one of the most important coenzymes involved in intracellular redox reactions including cancer metabolism,17 and it generally exists in two forms including oxidized NAD+ and reduced NADH.18 NAD+/NADH levels can affect cancer cell growth by inhibiting the related energy production pathways,6 such as glycolysis, the tricarboxylic acid cycle, and the mitochondrial electron transport chain (ETC). Hence, regulation of NAD+/NADH levels provides an effective strategy for the inhibition of cancer cells by driving the NAD+ and NADH interconversion.8,19,20 However, the regulated ratio of NAD+/NADH can be gradually restored to rebalance the redox levels through the cell repair process.11 Photocatalytic reactions exhibit great potential in achieving desirable intracellular reactions due to the strong controllability, mild reaction conditions,21 and noninvasive remote regulation.22,23 But the poor biocompatibility of noble metal catalysts24,25 could be harmful to living organisms resulting in limited applications in vivo. In comparison, water-soluble conjugated polymers (WSCPs) possess high biocompatibility26–29 and electron transfer efficiency that are beneficial to their applications in photoinduced processes in living organisms.30–32 The ability of WSCPs to catalyze cell-mediated polymerizations33–36 could be exploited in intracellular photocatalytic reactions, which create NAD+/NADH imbalance and regulate cell activities. In this work, we report an emerging photocatalytic reaction and realize the irreversible decomposition of NAD+ to nicotinamide and adenosine diphosphate (ADP)-ribose by the cationic poly(fluorene-co-phenylene) (PFP). The chemical reaction was induced in hypoxic tumor cells to successfully disrupt the cell activity (Scheme 1). The mechanistic investigation shows that the PFP acts as the photocatalyst in the system, and the photoelectrons transfer from the excited PFP (PFP*) to NAD+ to yield nicotinamide and ADP-ribose instead of the naturally occurring bioactive NADH. Therefore, the photocatalytic reaction mediated by PFP irreversibly regulates NAD+ rather than the common strategy based on a reversible reaction between NAD+ and active NADH.3 Inside these tumor cells, the reaction compromises mitochondrial functions that are mainly manifested as blockage of ETC, decreased mitochondrial membrane potential (MMP), and decreased adenosine triphosphate (ATP) level. The disturbed cellular redox cycle and irreversible damage of mitochondria by PFP under irradiation can cause apoptosis and finally disrupt the tumor cell activity. Scheme 1 | Schematic illustration of the photocatalysis reaction by PFP for irreversibly converting NAD+ to nicotinamide and ADP-ribose and inducing apoptosis of hypoxia cancer cells. Download figure Download PowerPoint Experimental Section General methods and materials Cationic PFP was synthesized according to the reported literature procedure.32 All chemical reagents are commercially available and used without further purification. 5,5-Dimethyl-1-pyrroline N-oxide (DMPO) was purchased from Sigma-Aldrich (St. Louis, Missouri, USA). 4T1 cells were purchased from the cell culture center of the Institute of Basic Medical Sciences, Chinese Academy of Medical Sciences (Beijing, China). Roswell Park Memorial Institute 1640 medium (RPMI 1640) was purchased from Hyclone (Beijing, China). Fetal bovine serum was purchased from Sijiqing Biological Engineering Materials (Hangzhou, China). 3-(4,5-Dimethylthiazol-2-yl)-2,5-diphenyltetrazolium bromide (MTT) was purchased from Xinjingke Biotechnology Co., Ltd. (Beijing, China). MMP assay kit (JC-1), ATP, and BCA Assay Kits were acquired from Beyotime Biotechnology (Shanghai, China). The NAD+/NADH Ratio Kit was purchased from AAT Bioquest (Pleasanton, California, USA). Deionized water was obtained from a Milli-Q system (Millipore, Bedford, MA, United States). 1,1′-dioctadecyl-3,3,3′,3′-tetramethylindodicarbocyanine (DiD) was provided by Life Technologies (Waltham, Massachusetts, USA). UV–vis absorption spectra were measured by a Thermo Scientific Evolution 201 spectrophotometer (Thermo Fisher, Waltham, Massachusetts, USA), and the fluorescence spectra were taken on a Hitachi F-4500 fluorometer (Hitachi, Tokyo, Japan) equipped with a Xenon lamp excitation source. Ultraviolet photoelectron spectroscopy (UPS) was conducted on an AXIS ULTRA DLD instrument (Kratos, Manchester, UK). Electrochemistry measurements were conducted on an electrochemical workstation (Autolab PGATAT 302N, Metrohm, Herisau, Switzerland). NMR spectra were obtained from a Bruker Avance HD 400 MHz and Bruker AVANCE 600 MHz (Rheinstetten, Germany). The electron paramagnetic resonance (EPR) spectra were measured by Bruker Xenoe nano (Bruker EMXnano, Fällanden, Switzerland). The MTT assay was recorded on a microplate reader (BioTek, Winooski, Vermont, USA). Confocal laser scanning microscopy (CLSM) images were conducted on confocal laser scanning biological microscope (Olympus FV 1200-BX61, Olympus, Hataya, Japan). Intracellular NAD+/NADH ratio assay 4T1 cells were treated in the medium at 37 °C at a density of 5 × 105 cells/well in 6-well plates overnight. Then, the medium with PFP (8 μM) and triethanolamine (TEOA; 4 mM) was used to cultivate 4T1 cells at 37 °C for 12 h. The cells were transferred to a hypoxic environment (1% O2, 5% CO2, 37 °C) for 6 h. After irradiation with 50 mW/cm2 for 20 min, the cells were treated with 250 μL cell lysis buffer. The centrifugal supernatant was collected and used to measure intracellular NAD+/NADH ratio by commercial kit. MMP measurement The 4T1 were added in confocal dishes at a density of 8 × 104 cells/mL and treated in the RPMI medium containing PFP (8 μM) and TEOA (4 mM) at 37 °C for 12 h. Then the cells were incubated in 1% O2 and 5% CO2 at 37 °C for 6 h. After irradiation under white light with 50 mW/cm2 for 30 min, the 4T1 cells were incubated with JC-1 (5×) for 20 min. Then cells were rapidly washed three times by JC-1 (1.25×). The CLSM images were collected at 520–550 nm (λex, 488 nm), and 575–675 nm (λex, 488 nm), respectively. EPR A mixture of 100 μL PFP (1 mM, dimethyl sulfoxide), 20 μL TEOA (1 M), 100 μL DMPO (1 M), 200 μL NAD+ (50 mM), and 580 μL phosphate-buffered saline was irradiated under white light (50 mW/cm2), and the EPR spectra were recorded every 5 min. (Center of Field, 3443.90 G; Sweep Width, 200.0 G; Power, 5 mW; Power Attenuation, 20.00 dB; Modulation Amplitude, 1.0 G; Number of Scans, 6.) Results and Discussion Characteristics of photocatalyst To verify the feasibility of photocatalytic reaction of PFP to catalytically convert NAD+, the photophysical information, energy level, and fluorescence decay kinetics of PFP were collected accordingly. As shown in Figure 1a, PFP exhibited a maximum absorption wavelength of 380 nm, and the peak of fluorescence emission was 429 nm. The band gap of PFP was calculated with a standard method to be 2.97 eV.4 The photoelectric response under a solar simulator demonstrated the possible conversion of absorbed photons into excited electrons ( Supporting Information Figure S1). The highest occupied molecular orbital (HOMO) was determined to be −5.82 eV by UPS (Figure 1b), and the lowest unoccupied molecular orbital (LUMO) was calculated to be −2.85 eV with an optical band gap of 2.97 eV. The Ered and Eox of PFP were +1.32 V and −1.65 V (vs normal hydrogen electrode [NHE]), respectively, obtained by calculations described in the literature.37,38 The reduction potential of NAD+/NADH is −0.54 V versus NHE,39 which is lower than Ered of PFP (+1.32 V vs NHE), and, therefore, suitable to capture the electron from PFP*. The fluorescence decay kinetic curves show that the fluorescence lifetime of PFP decreased from 104 to 23 ns with the addition of NAD+ (Figure 1c). The electron-transfer pathway from PFP to NAD+ under illumination was predicted based on the HOMO/LUMO potential (Figure 1d). The electron captured from the HOMO of PFP could be transferred to NAD+ in the reaction process. All the above characterizations indicate that under illumination PFP generates the holes and electrons, and then the photoelectrons from the PFP HOMO to LUMO transition might reduce NAD+ into the product. The holes are consumed by TEOA whose redox potential (+0.53 V vs NHE) is more negative than PFP (+1.32 V vs NHE). Figure 1 | Material property characterizations of conjugated polymer PFP. (a) Normalized absorption and fluorescence spectrum (in H2O at 25 °C). (b) Ultraviolet photoelectron spectrum of PFP. (c) The fluorescence decay kinetic curves of PFP (10 μM) and PFP (10 μM) + NAD+ (1 mM) with 100 mM TEOA in H2O at 25 °C (λex: 405 nm). (d) The redox potential of the elements involved in the reaction pathway. Download figure Download PowerPoint The photocatalysis reaction to decompose NAD+ According to NMR and mass spectrometry, the photocatalytic reaction of NAD+ generated nicotinamide and ADP-ribose as the products (Figure 2a and Supporting Information Figure S2). Before illumination, 1H NMR peaks corresponding to the nicotinamide unit (a–d), adenine unit (g and h), and the two glycosidic protons (e and f) in NAD+ were identified (black trace), whereas PFP peaks were invisible due to the low concentration. After illumination, the complicated 1H NMR spectral pattern consisted of free nicotinamide (a′–d′, red trace), ADP-ribose (e′–h′), and unreacted NAD+ (not labeled). Major changes of the peaks of nicotinamide and the glycosidic proton e/e′ indicate that the photocatalytic cleavage occurred between nicotinamide and its adjacent ribose. In contrast, minor peak shifts of adenine and its adjacent glycosidic proton f/f′ suggest an intact glycoside on adenine. To further resolve the products, diffusion ordered spectroscopy (DOSY) was performed on the sample after 1 h illumination (Figure 2b). Peaks corresponding to the free nicotinamide exhibited two- to threefold higher diffusion coefficients than other major peaks, and the trend was consistent with the smaller molar mass of nicotinamide (122 g/mol) compared to NAD+ and ADP-ribose. Although the latter two compounds could not be resolved in 1H NMR or DOSY spectra, they were identified on the electrospray ionization-mass spectrometry (ESI-MS) spectrum. The peaks at m/z of 558.1 and 540.1 correspond to ADP-ribose and its dehydrated derivatives, respectively, (Figure 2c), and the peak at 662.1 is unreacted NAD+. To monitor the reaction progress to characterize the kinetics of the photocatalysis, the 1H NMR integrations of H(d) in NAD+ and H(d′) in nicotinamide were calculated (Figure 2d). After 1 h irradiation, 113 μM nicotinamide was obtained, and the products continued to accumulate with extended irradiation time. Figure 2 | The reaction of NAD+ with PFP as the photocatalyst. (a) 1H NMR (D2O) spectra of the reaction mixture before (black) and after illumination (red). (b) DOSY spectrum for further resolving nicotinamide in the illuminated reaction mixture. (c) Negative ion ESI-MS indicates photocatalytic generation of ADP-ribose. (d) Nicotinamide generation under an extended time of illumination. Download figure Download PowerPoint Intracellular catalysis study and therapeutic efficacy The intracellular catalytic effect of PFP photocatalyst on 4T1 cancer cells was investigated. The stability of the cationic conjugated PFP in various physiological solutions was measured ( Supporting Information Figure S3). The similar absorption spectra indicate excellent stability of the conjugated PFP skeleton as a photocatalyst. Meanwhile, the efficiency of this photocatalytic reaction was measured under hypoxic and acidic conditions to simulate the tumor microenvironment. As shown in Supporting Information Figure S4, the efficiency of the photocatalytic reaction was not affected by the low oxygen concentration or acidic pH. It indicates that this photocatalytic reaction mediated by PFP worked in the tumor microenvironment. To exclude the photosensitization effects of PFP, the reactive oxygen species (ROS) production capacity of PFP was measured by 2,7-dichlorodihydrofluorescein (DCFH; Supporting Information Figure S5). In contrast to the efficient photogeneration of ROS under aerobic conditions, negligible ROS were produced by the illumination of PFP under hypoxic conditions. It indicates that the influence of common ROS is not a major contributing factor in this antitumor system. As illustrated in Supporting Information Figure S6, CLSM images of 4T1 cells showed that the blue signal from PFP was surrounded by an entire membrane (labeled with DiD and emitting red signal), which elucidates the successful internalization of PFP. As shown in Supporting Information Figure S7, the PFP, which located in the lysosome, was taken up into cells over the incubation time and retained intracellularly for more than 72 h. This indicates that PFP has the potential to act as a long-term catalyst in cancer cells. To verify that the catalytic reaction could occur in cells, the intracellular NAD+/NADH ratio in hypoxic 4T1 cells treated with PFP under light irradiation was measured. As shown in Figure 3a, the NAD+/NADH ratio of the optical irradiation group treated with PFP decreased from 73% to 65% compared to the dark group. Thus, PFP can catalytically convert NAD+ into nicotinamide and ADP-ribose under white illumination. As an important coenzyme pair in the mitochondrial ETC, and a raw material for photocatalytic reactions, NAD+ can affect the function of mitochondria. The continuous depletion of NAD+ can lead to mitochondrial apoptosis, thereby affecting cell growth. So, the related MMP was measured to further investigate the influence of the explored photocatalytic reaction in hypoxic 4T1 cells. The commercial MMP probe JC-1 was used to visually observe the apoptosis of mitochondria after treatment with PFP and optical illumination. Typically, JC-1 characterizes the state of MMP (ΔΨm) by its monomer or J-aggregate. The J-aggregate initially attaches to the outer membrane of the normal mitochondria and emits red fluorescence. However, the ΔΨm decreases once the NADH-dependent oxidative phosphorylation (OXPHOS) process is inhibited,40 and the JC-1 changes to the monomer and shows green fluorescence, indicating the mitochondria goes into apoptosis (Figure 3b). As displayed in Figure 3c and Supporting Information Figure S8, 4T1 cells exhibited strong green fluorescence after being treated with PFP upon optical illumination in the hypoxic environment, suggesting that the ΔΨm decreased and the mitochondria may be undergoing apoptosis. The variations of the intracellular ATP level were subsequently measured to confirm the mitochondria damage. As shown in Figure 3d, the ATP level of 4T1 cells in the control group had negligible changes no matter with or without white-light illumination. However, the ATP level of the white-irradiation group decreased to 0.11 μg/mg protein (from 0.20 μg/mg protein), which indicates that the process of NAD+ cleavage and product production led to mitochondrial damage, further affecting the normal metabolic process and finally causing cell apoptosis. To visually show the therapeutic effect of the PFP system on 4T1 cancer cells, LIVE/DEAD cell staining was performed accordingly. As shown in Figure 3e and Supporting Information Figure S9, almost all the cells treated with PFP in the dark emitted strong green signals of living cells, indicating a high cell-survival rate and the promising biosafety of PFP, whereas over half of the cells treated with PFP under illumination showed the red signal indicative of dead cells, signifying that PFP had an obvious therapeutic effect on the life activities of tumor cells under white-light illumination. The MTT assay was performed to evaluate the cell viability in the presence of different concentrations of PFP. 4T1 cells had high cell viability even treated with a high concentration of PFP (16 μM) in dark conditions (Figure 3f), further confirming its biocompatibility. However, upon white-light illumination, the 4T1 cell viability apparently decreased as the concentration of PFP increased (over 2 μM). 4T1 cell viability fell below 60% after being treated with 16 μM PFP, reinforcing the good therapeutic efficacy of the artificial photocatalytic system due to the catalytic decomposition of NAD+ by this system that results in a disordered metabolic process. Electron sacrificing agent TEOA and the nicotinamide involved in the system also exhibited negligible toxicity ( Supporting Information Figures S10 and S11). As illustrated in Figure 3g and Supporting Information Figure S12, the apoptosis cell ratio of 4T1 cells after being treated with PFP under white-light illumination increased significantly from 0.5% to 42.4%, whereas the control groups remained negligibly or slightly changed, including the white-light illumination-only group and PFP-only group. This indicates the good therapeutic efficacy of the artificial photocatalytic system as well as the noninvasiveness and biocompatibility of optical operation manner and PFP photocatalyst, respectively. To study the universality of the described photocatalytic system, two other tumor cell lines, HeLa and HepG2, were characterized by MTT assay ( Supporting Information Figure S13). With increasing concentration of PFP (over 1 μM), the decreasing cell viability of HeLa and HepG2 demonstrate that the photocatalytic system was effective for various cancer cells, including human cancer cells. Compared to the common strategy of regulation between NAD+ and active NADH, the metabolism regulation catalyzed by PFP is an irreversible reaction, which can disturb the cellular redox cycle and promote irreversible damage to mitochondria and even cells. The cytotoxicity of the photocatalytic reaction to human foreskin fibroblasts (HFF) cells was measured to study the effect of this photocatalytic reaction on normal cells ( Supporting Information Figure S14). The HFF cells maintained high viability under light and dark conditions. Figure 3 | Evaluation of the cell apoptosis by PFP photocatalyst upon optical illumination. (a) Changes of the intracellular NAD+ concentration catalyzed by PFP under optical illumination for 30 min. (b) Schematic illustration of measuring mitochondrial membrane potential changes by MitoProbe JC-1. (c) Confocal laser scanning microscopy images to show mitochondrial membrane potential (ΔΨm) changes of 4T1 cells after incubation with PFP with or without optical irradiation. The fluorescence imaging of JC-1 was collected at 575–675 nm (λex, 559 nm), 520–550 nm (λex, 488 nm), respectively. Scar bar, 50 μm. (d) The intracellular ATP levels before and after catalysis by PFP under optical illumination for 30 min. (e) Staining of living and dead cells by calcein (AM) and propidium iodide (PI). The fluorescence imaging of AM and PI were collected at 520–565 nm (λex, 488 nm), 575–675 nm (λex, 559 nm), respectively. Scar bar, 200 μm. (f) Cell viability of 4T1 cells after being treated with PFP (0.25–16 μM) with or without white-light illumination (50 mW/cm2) for 30 min. (g) Apoptosis and necrosis of 4T1 cells after different treatments were analyzed using flow cytometry according to the fluorescence intensity in fluorescein isothiocyanate (FITC) and propidium iodide (PI) channels. Download figure Download PowerPoint The radical and biological action characterizations of the photocatalysis reaction EPR assay was conducted to reveal the specific mechanism of the above photocatalytic process. Because the transient radical would be trapped by DMPO and changed to the EPR-active species DMPO-ADP-ribosyl•, the collected carbon-free radical signal under irradiation was recognized as ADP-ribosyl• radical (Figure 4a). It demonstrates that the NAD+ transferred to ADP-ribosyl• radical in this photocatalysis process under illumination. Regarding the discussion of Figure 1d, the electron that NAD+ captured was from PFP*. Based on the above experimental results, a presumptive mechanism of photocatalytic reaction by PFP is proposed (Figure 4b). Under illumination, electrons from PFP* transfer to NAD+, generating ADP-ribose• radical and nicotinamide; then the holes are captured by TEOA. After electron rearrangement, ADP-ribose is obtained from the NAD+· by the combination of water. To test the irreversibility of this system in vivo, the enzymatic activity of production was characterized by monitoring the catalytic reaction of pyruvic acid with lactic dehydrogenase (LDH). Generally, LDH catalyzes the conversion of pyruvic acid to lactic acid with the oxidation of NADH, and the reaction should be inhibited when NADH is replaced by photocatalytic products. The enzymatic activity of production was characterized by monitoring the absorption at 340 nm, which is the characteristic absorption of NADH. As shown in Figure 4c, the absorption at 340 nm of photocatalytic products exhibited negligible change with the addition of the pyruvic acid and LDH, demonstrating that the photocatalytic product could not be used as a coenzyme of LDH. For active NADH, the absorption decreased significantly after adding the pyruvic acid and LDH. Thus, photocatalytic products are unable to participate in the normal enzymatic reactions, which proves that the products obtained in the photocatalytic reaction can not re-enter the enzymatic cycle of the cells. Therefore, the photocatalytic reaction interferes with the function of cancer cells and induces hypoxic tumor apoptosis. Figure 4 | The radical and biological action characterizations of products from NAD+ catalyzed by PFP under light illumination. (a) EPR spectrum of carbon radical trapped by DMPO under irradiation (white light, 50 mW/cm2). (b) The mechanism of the photocatalytic reaction by PFP for irreversibly converting NAD+ to nicotinamide and ADP-ribose. (c) The absorption of the PFP + NAD+ group with LDH under dark and light, compared to the NADH group. Biocatalysis upon production under irradiation compared with NADH catalyzed by LDH. Download figure Download PowerPoint Conclusions In summary, we report a new photocatalytic reaction where irradiated PFP irreversibly converts NAD+ to nicotinamide and ADP-ribose. The chemical reaction was