Oxidative photocatalysis on membranes triggers non-canonical pyroptosis

doi:10.21203/rs.3.rs-3126446/v1

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Oxidative photocatalysis on membranes triggers non-canonical pyroptosis

https://doi.org/10.21203/rs.3.rs-3126446/v1

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published 13 May, 2024

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Intracellular membranes composing organelles of eukaryotes include membrane proteins playing crucial roles in physiological functions. However, a comprehensive understanding of the cellular responses triggered by intracellular membrane-targeted oxidative stress remains elusive. Herein, we developed an amphiphilic photocatalyst localised in intracellular membranes to damage membrane proteins oxidatively, resulting in non-canonical pyroptosis. Our developed photocatalysis generated hydroxyl radicals and hydrogen peroxides via water oxidation, which was accelerated under hypoxia. Single-molecule magnetic tweezers revealed that photocatalysis-induced oxidation markedly destabilised membrane protein folding. In cell environment, label-free quantification revealed that oxidative damage occurred primarily in membrane proteins related to protein quality control, thereby aggravating mitochondrial and endoplasmic reticulum stress and inducing lytic cell death. Notably, the photocatalysis activated non-canonical inflammasome caspases, resulting in gasdermin D cleavage to its pore-forming fragment and subsequent pyroptosis. These findings suggest that the oxidation of intracellular membrane proteins triggers non-canonical pyroptosis.

Physical sciences/Chemistry/Chemical biology/Chemical tools

Biological sciences/Biochemistry/Chemical modification

Biological sciences/Cell biology/Cell death

The intracellular membrane in eukaryotes serves as a reaction platform for biochemical processes, such as organelle interactions, signalling, metabolic reactions, and biomolecular productions^1-3. These physiological reactions are mediated by membrane proteins and lipids³, which are the functional constituents of the intracellular membranes. However, these components are susceptible to oxidative damage caused by the endo- and exogenous reactive oxygen species (ROS)^4,5. This oxidative damage to the intracellular membrane can disrupt the organellar function and trigger programmed cell death, which is implicated in the pathogenesis of various diseases^6-8. Discovering cellular processes in response to oxidative stress on intracellular membranes is essential for devising strategies to control cell death signalling and treat diseases related to membrane protein oxidation. It has been reported that lipids constituting intracellular membranes can be peroxidised by radical propagation reactions through cytosolic iron-induced hydroxyl radical (•OH) generation, which eventually causes caspase-independent ferroptosis⁹. However, cell death signalling in response to the intracellular membrane-localised ROS generation and oxidative stress on intracellular membrane proteins have not been fully elucidated.

Oxidative photocatalysis can be a promising approach to control oxidative stress at desired points inside cells spatiotemporally¹⁰. Recent studies have investigated cellular responses to organelle-targeted oxidative stress and subsequent cell death signalling using the photocatalysts generating ROS at specific organelles^11,12. However, a photocatalyst capable of simultaneous localization within the intracellular membranes enclosing organelles (such as the endoplasmic reticulum, Golgi apparatus, mitochondria, vesicles, and nucleus) has not been developed. This limitation hinders the exploration of cellular responses to membrane protein oxidation within the intracellular membranes. Nevertheless, the development of an intracellular membrane-targeted photocatalyst presents a significant challenge, as these photocatalysts should possess high lipophilicity to localise within the intracellular membrane while still being able to penetrate the plasma membrane. Therefore, developing intracellular membrane-targeted oxidative photocatalysis is essential for analysing cellular responses to intracellular membrane oxidation.

Herein, we develop an intracellular membrane-localised organic photocatalyst, BTP, a fatty acid-like molecule consisting of hydrophobic linear π-conjugation and a hydrophilic head. Owing to its amphiphilic structure, BTP was localised in the intracellular membrane. Furthermore, its photocatalysis oxidised water molecules, resulting in the generation of highly oxidising radical species (i.e., hydroxyl radicals) and induction of intracellular membrane-targeted oxidative stress even under hypoxia. This oxidative damage irreversibly disrupted the folding stability of membrane proteins. As a result of the oxidative photocatalysis on intracellular membranes, we found that photocatalysis-induced oxidation occurred mainly in membrane proteins related to protein quality control (PQC), leading to the accumulation of misfolded proteins.

Notably, we initially found that the intracellular membrane oxidation activated non-canonical inflammasome caspases (4 and 5) and triggered subsequent gasdermin-D (GSDMD)-driven pyroptosis, which is an immunogenic and inflammatory cell death process mediated by inflammasomes.^13–15 Considering that pyroptosis of cancer cells can stimulate immune responses against tumours,^16–18 identifying chemical stimuli capable of inducing pyroptosis and elucidating their underlying mechanisms hold promise for providing strategies to promote antitumour responses. To this end, these results provide a potential approach to induce the activation of non-canonical inflammasome caspases and consequent pyroptosis through intracellular membrane-targeted oxidation.

Photocatalytic cycles to generate oxidising radical species

We synthesised BTP by pairing benzothiadiazole and triphenylamine, which are an electron donor and acceptor, respectively. This donor-acceptor type molecular structure promotes the charge separation ability of its excitons, thereby enhancing its electron transfer activity as a photocatalyst. We measured the photophysical and electrophysical properties of BTP to estimate its redox potentials of BTP (Extended Data Fig. 1a–c). Based on the ground and excited state redox potentials of BTP, we suggest that the photocatalytic cycle of BTP causes membrane oxidation (Fig. 1b).

Under light irradiation, the excited state of BTP (BTP*) exists for a few nanoseconds, generating E*^(0/−) and E*^(+/0)which are the reduction and oxidation potentials of BTP*, respectively. In aqueous environments, only neighbouring molecules, such as water and oxygen, can allow for the electron transfer from water to BTP* (E*^(0/−)) and from BTP* to oxygen (E*^(+/0)). Each redox potential was estimated using cyclic voltammetry and band gap energy (see Methods). The E*^(0/−)of BTP was 1.47 V (vs. the normal hydrogen electrode), which enables water oxidation via a two-electron pathway yielding H₂O₂(E^(H₂O₂/^H^₂^O⁾ = 1.34 V at pH 7; Fig. 1b, Extended Data Fig. 1d ①)^19-21. To examine the electron transfer between water and BTP*, a fluorescence quenching assay was performed (Fig. 1c). The BTP fluorescence gradually decreased as the water content of acetonitrile solution increased and was mostly quenched at 10% water content owing to the reductive electron transfer from water to BTP*. Furthermore, the excited lifetime of BTP* was diminished as water content increased (Extended Data Fig. 1e). These results imply that BTP* was reductively quenched via water oxidation.

Correspondingly, we assayed H₂O₂, the expected product of water oxidation, using peroxidase and N,N-diethyl-p-phenylenediamine (DPD)²² (Fig. 1d). BTP photoexcitation generated H₂O₂ in aqueous solution, whereas BTP* could not produce H₂O₂ without water (in dimethyl sulfoxide). This result revealed that H₂O₂ was generated from water during BTP photocatalysis. Furthermore, we found that hypoxia substantially accelerated photocatalytic H₂O₂ production, implying that oxygen functions as quencher of BTP* via oxygen reduction (E*^(+/0) = −1.36 V, E^{(O₂/O₂•−)} = −0.33 V; Fig. 1b, Extended Data Fig. 1d ③)²³. The oxygen reduction reaction competes with water oxidation by BTP*, indicating that hypoxia could enhance the H₂O₂ generation. When BTP* accepts an electron, it transforms into BTP^•⁻.BTP^•− can easily donate its electron to a nearby H₂O₂ propagating to a hydroxyl radical (•OH; E^(0/−) = −0.89 V, E^{(H₂O₂/•OH)} = 0.38 V; Fig. 1b, Extended Data Fig. 1d ②)²⁴. Therefore, we performed a hydroxyphenyl fluorescein (HPF) assay²⁵ to confirm •OH generation (Fig. 1e, Extended Data Fig. 1f). The HPF assay revealed that BTP photocatalysis generated •OH, which was promoted under hypoxic and H₂O₂-supplemented conditions. This result implies that •OH is generated from H₂O₂, and this reaction is escalated under hypoxia, considering that oxygen functions as an inhibitor of H₂O₂ generation by quenching BTP*. Furthermore, •OH production was impaired in the absence of water (in dimethylformamide), implying that •OH is also produced from water-mediated photocatalysis (Fig. 1e). Electron paramagnetic resonance (EPR) spectroscopy with 5-tert-butoxycarbonyl-5-methyl-1-pyrroline N-oxide (BMPO, spin trap for •OH)²⁶ followed to clarify •OH generation (Fig. 1f).

When oxygen is reduced by BTP*, superoxide radicals (O₂^•−) can also be produced, and BTP^•+ can accept an electron from nearby amino acids, such as Trp, Tyr, and Cys (see Supplementary Information; Fig. 1b, Extended Data Fig. 1d ④, g–I). BTP photocatalysis generates H₂O₂ and •OH but does not produce singlet oxygen (¹O₂; Extended Data Fig. 1j). •OH is highly oxidising reactive oxygen species (ROS) that is sufficient for membrane component oxidation^27-29. Therefore, we investigated the oxidation of methionine and unsaturated lipids. High-resolution mass spectrometry (HR-MS) revealed the oxidised products of Met and phosphatidylcholine (Extended Data Fig. 1k–l), further exhibiting the potential to inflict severe oxidative stress to the cellular membrane.

Oxidative damage on membranes by BTP photocatalysis

We next examined where BTP is located and oxidative stress is produced in cells. Co-localisation experiments revealed that BTP was in the endomembrane system, including Golgi apparatus (GA) and ER, but not in mitochondrial membranes (Extended Data Fig. 2a). However, BTP photocatalysis changed the localisation pattern from the ER membrane to the mitochondrial membrane (Extended Data Fig. 2b), as confirmed by structured illumination microscopy (SIM) with viable HeLa cells (Extended Data Fig. 2c). This change in location is likely because BTP photocatalysis reduces the ER membrane integrity³⁰, leading to its migration to nearby membranes. Furthermore, we found that BTP photocatalysis generated substantial membrane oxidative stress. The dichlorodihydrofluorescein diacetate (DCFH₂-DA, a ROS indicator) assay showed that BTP photocatalysis increased DCF fluorescence in HeLa cells (Extended Data Fig. 2d, e), and the lipid peroxidation assay with BODIPY^TM 581/591 C11 showed that lipid peroxides were generated only in the light-irradiated region (Extended Data Fig. 2f, g). Additionally, we established BTP photocatalysis-induced generation of O₂^•−using a dihydroethidium (an O₂^•− sensor) assay (Extended Data Fig. 2h). These results indicate that BTP photocatalysis induces oxidative stress and lipid peroxidation on intracellular membranes.

Destabilisation of membrane protein fold by BTP photocatalysis

Membrane proteins are crucial components of intracellular membranes, and their structural damage leads to impaired functions, ultimately affecting cell fate and programmed death signalling³¹. Thus, using an in vitro membrane protein stability assay (Fig. 2a–d), we examined the effects of BTP photocatalysis on membrane protein folding. We adopted E. coli rhomboid protease GlpG, a widely used model protein for studies on membrane protein folding and stability^32-34. Only under the condition of BTP added and blue light exposure (30 J·cm⁻²), the intensity of the gel band for GlpG (~22 kDa) was decreased to 31±9.0% from the negative control at 150 μM BTP (Fig. 2a, b), whereas the band intensities for GlpG aggregates (>23 kDa) were increased (Fig. 2a). This indicates the destabilisation and aggregation of GlpG oxidatively damaged by BTP photocatalysis. This result was also supported by thermal denaturation assay (Fig. 2c, d). Indeed, the resistance to the thermal denaturation of GlpG was reduced by the oxidative damage. The transition midpoint temperature of the thermal denaturation was largely decreased from 75 ℃ at no BTP condition to 35 ℃ at 150 μM BTP (Fig. 2c, d).

To confirm the oxidative destabilisation of GlpG by which protein aggregation was promoted, we used a single-molecule tweezer approach that could entirely exclude the aggregation events (Fig. 2e–h)^32,35-37. In this approach, a single GlpG embedded in a lipid bilayer disc (bicelle) was tethered via DNA handles to a magnetic bead and the sample chamber surface (Fig. 2e)^37,38. Using a pair of magnets generating the magnetic field gradient, we applied a mechanical force of a few to tens of pN to the tethered GlpG. The repetitive unfolding was reproducible by more than a hundred cycles under the condition of blue light exposure (λ = 450 nm, 9.16 mW·cm^–²) without BTP (Fig. 2f, upper), and the unfolding forces were distributed at 34±5.8 pN (Fig. 2g; Extended Data Fig. 3a for infrared light of λ = 850 nm, 39.51 mW·cm^–²). Upon the addition of BTP with blue light exposure, however, the unfolding forces were drastically reduced to less than 15 pN at a critical point in the pulling cycle (C₀), even unmeasurable at 20 μM BTP (Fig. 2f, middle and lower). After the C₀ point, the unfolding force values were not restored to that of the normal level for the GlpG unfolding (Fig. 2f, g). The transition occurred in an earlier pulling cycle (earlier time) at a higher BTP concentration (Fig. 2h). Moreover, the injection of fresh bicelles after C₀ was unable to restore the normal unfolding, indicating that the major effect was on the membrane protein, not the lipid bilayer environment (Extended Data Fig. 3b). These results imply that, once photocatalytic oxidative damage is made to the single membrane protein, the interactions in native protein fold are drastically destabilised.

Impact of BTP photocatalysis on protein quality control proposed by proteomics

To identify the membrane proteins oxidised by BTP photocatalysis in the cells, we analysed the oxidative modifications of membrane proteins. Because methionine is one of the most labile amino acids under oxidative stress, proteins comprising of oxidised methionine residues (O-Met) were analysed using label-free quantitative mass spectrometry (Extended Data Fig. 4a)^28,39,40. Triplicate experiments detected a total of 5143 proteins and 805 “significantly oxidised proteins” as defined by a p-value < 0.1 and fold change > 2, following BTP photocatalysis of HeLa cells (Fig. 3a). The oxidised proteome was then classified by cellular localisation and function (Fig. 3a–d, Extended Data Fig. 5). Remarkably, the significantly oxidised proportion of membrane proteins was 1.7 times higher than that of non-membrane proteins (22.4% and 13.1%, respectively; Extended Data Fig. 4b). This result aligns with the membrane-anchoring property of BTP. Additionally, the proportions of significantly oxidised membrane proteins of the ER, GA, and mitochondria were 22.4%, 15.9%, and 35.4%, respectively (vs. detected membrane proteins of each organelle; Extended Data Fig. 4c). This proportion is much higher than that of lumen proteins (11.7%), implying that BTP photocatalysis oxidises the nearby membrane proteins of the ER, GA, and mitochondria.

Notably, the ER, GA, and mitochondria, three organelles crucial for protein quality control (PQC)⁴¹, possessed 71.1% of the significantly oxidised membrane proteins resulting from BTP photocatalysis (Fig. 3c). We therefore focused on PQC-related functions, including the unfolded protein response (UPR), translation, and protein transport. Among 313 significantly oxidised membrane proteins, 84 proteins were clustered into four functional networks, i) UPR, ii) mitochondrial translation, iii) ER–GA transport, and iv) lipid metabolism (Fig. 3d, Extended Data Fig. 6). Gene Ontology (GO) functional annotations concerning biological processes, such as transport, translation, and localisation-related proteins, were found in the O-Met proteome (Fig. 3e). The O-Met proteome involves proteins responsible for ER-related degradation and UPR modulation, such as SYVN1, ERLIN2, ERO1L, STT3A, and Rab6B (Fig. 3f, Extended Data Fig. 4d). Simultaneously, the oxidation of mitoribosomal proteins was identified in the O-Met proteome, including MRPS, MRPL families, and MTG1. Furthermore, ER antero/retrograde transport proteins, such as YIF1A, TMEM10 and Rab21, and the proteins responsible for lipid metabolism, STS, POR, and HSD17B7, were significantly oxidised by BTP photocatalysis. As described in the folding stability experiments (Fig. 2), the significantly oxidised membrane proteins might have lost their folding stability. Therefore, we hypothesised that BTP-induced oxidation and the resulting dysfunction of proteins could deteriorate PQC and escalate UPR stress.

Cation mobilisation by BTP photocatalysis

BTP photocatalysis causes the destabilisation of the membrane protein structure related to PQC, leading to an irreversible accumulation of misfolded proteins, thereby enhancing stress on the ER, GA, and mitochondria^42,43. Failure of the PQC and subsequent maladaptive UPR has been reported to trigger Ca²⁺ mobilisation⁴⁴, and we confirmed this using Rhod-2 (a Ca²⁺ indicator). Ca²⁺ concentration in the mitochondria increased considerably following light exposure (0.3 mW) for 30 s, indicating mitochondrial Ca²⁺ uptake (Fig. 4a). The line-cut analysis supported that the MitoTracker signal was well merged with the increased Rhod-2 signal (Fig. 4b, c) after BTP photocatalysis. This tendency was also observed in flowcytometry with Rhod-2 (Fig. 4d). Accumulation of misfolded proteins and Ca²⁺ leads to osmotic swelling, resulting in mitochondrial dysfunction. We observed BTP photocatalysis-induced mitochondrial swelling accompanied by fission and fusion, known as the mitochondrial PQC process (Fig. 4e)⁴⁵. Simultaneously, we found intracellular K⁺ efflux after BTP photocatalysis using flowcytometry with ION K⁺ Green-2 (K⁺indicator; Fig. 4f). These results show that oxidative photocatalysis induces maladaptive UPR and cation mobilisation in response to oxidative damage of intracellular membrane.

Non-canonical inflammasome caspases-induced pyroptosis

Given the impact of BTP photocatalysis on membranes, we examined how BTP photocatalysis might impact cell death signalling responses. HeLa cell viability was tested by examining propidium iodide (PI) or calcein AM uptake, and the MTT assay. The results showed that nearly all HeLa cells died within 24 hours following BTP photocatalysis (Extended Data Fig. 7a, b). Even in hypoxic pancreatic cancer cell lines (Panc-1 and MiaPaca-2), MTT assays indicated severe toxicity caused by escalated ∙OH generation under hypoxia (Extended Data Fig. 7c). An examination of the cellular morphology indicated that BTP photocatalysis caused a lytic-type cell death, consistent with a pyroptotic morphology; including plasma membrane swelling and abnormal blebbing (Fig. 5a)^13,15. BTP photocatalysis induced lactate dehydrogenase (LDH) release, a measure of plasma membrane rupture, to a greater extent than the pyroptotic stimuli, LPS and nigericin, or the photosensitiser (Ce6) used in photodynamic therapy (Fig. 5b), indicating that cell death induced by BTP photocatalysis is highly lytic. Contrary to apoptosis, PI penetrated the plasma membrane but not the nuclear envelope within an hour after BTP photocatalysis (Extended Data Fig. 7d), implying that the nuclear envelope collapses only after plasma membrane integrity is compromised. Based on these cell death characteristics, we hypothesised that BTP photocatalysis triggers pyroptosis.

In many cases, ∙OH production and lipid peroxidation are associated with ferroptosis; thus, we first distinguished the characteristics of cell death caused by BTP photocatalysis from ferroptosis. Liproxstatin-1 (a lipid peroxidation and ferroptosis inhibitor) and z-VAD-fmk (a pan-caspase inhibitor) were used to investigate whether caspases or lipid peroxidation are involved in the lytic cell death triggered by BTP photocatalysis. Interestingly, z-VAD-fmk treatment reduced LDH release caused by BTP photocatalysis, while Liproxstatin-1 was less effective (Extended Data Fig. 7e). In addition, substantial levels of membrane blebbing and PI penetration were still observed in Liproxstatin-1-treated cells, but not in z-VAD-fmk-treated cells (Extended Data Fig. 7f). These results indicate that BTP photocatalysis-triggered cell death depends on caspase activation rather than on lipid peroxidation.

GSDMD cleavage by inflammatory caspases (i.e., caspase-1, -4, or -5) releases the N-terminal domain (GSDMD-NT), which then enacts pyroptosis via forming pores in the plasma membrane, resulting in non-selective ionic flux and the release of cellular immunogenic molecules¹⁴. Notably, BTP photocatalysis resulted in increased detection of GSDMD-NT in the cell lysates and cell media (Fig. 5c). Furthermore, pyroptotic morphology and LDH release caused by BTP were eliminated in GSDMD^-/- iBMDMs when compared to wildtype (WT) control macrophages (Fig. 5d, e). Because GSDMD is typically cleaved by caspase-1 that is engaged by canonical inflammasomes¹⁴, or by caspase-4/5, referred to as non-canonical inflammasomes⁴⁶, we examined the processing-associated activation of these caspases using western blot. Unexpectedly, these results showed that BTP photocatalysis causes cleavage of caspase-4/5 rather than caspase-1 (Fig. 5f). We further confirmed ATP secretion by caspase-4/5 and GSDMD activation (Fig. 5g), while the efficient secretions of cleaved interleukins (IL-18 and IL-1β) were not observed in the western blot analysis due to the lack of active caspase-1 (Fig. 5f). Considering that ER stress or cation mobilisation has been implicated in the activation of caspase-4/5^47-50, we surmise that the accumulation of misfolded proteins by BTP photocatalysis may trigger the cleavage of caspase-4/5 and subsequent pyroptosis.

We propose that photocatalytic membrane oxidation triggers non-canonical pyroptosis using the amphiphilic organic photocatalyst, BTP (Extended Data Fig. 8). Via photocatalysis, BTP generates highly oxidising •OH in a spatiotemporally controlled manner even under hypoxic conditions, thereby damaging the structural stability of membrane proteins. The single-molecule tweezer approach verified that BTP photocatalysis disrupts membrane protein folding. Using the methionine-oxidised proteome from the label-free quantification, we found that BTP photocatalysis substantially oxidised PQC-related membrane proteins of ER, GA, and mitochondria in cells. Disruption of the folding stability and oxidation of PQC-related proteins stimulate the accumulation of misfolded proteins, followed by ER stress, maladaptive UPR, and cation mobilisation. These cellular responses trigger the caspase-4/5-induced GSDMD cleavage and subsequent pyroptosis. In conclusion, we developed a chemical strategy to trigger non-canonical pyroptosis by photocatalytic membrane oxidation. This method can be widely used to spatiotemporally induce caspase-4/5 activation and pyroptosis in pathogenesis studies and clinical applications, even under hypoxic conditions.

The methods for organic synthesis of photocatalysts, characterisation, photophysical property analysis (UV-vis, photoluminescence spectroscopy, and time-correlated single photon counting), ROS assays (ABDA, H₂DCF-DA, and DHE assays), photocatalytic amino acid and lipid oxidation, laser scanning microscopy (LSM) and structured illumination microscopy (SIM) imaging, detailed preparation for LC-MS/MS, cation mobilisation assays, cell viability tests, and all experiments excluded from this section are provided in the Supplementary Information.

Cyclic voltammetry.

Cyclic voltammetry (CV) was conducted using a Vertex Potentiostat/Galvanostat (IVIUM Technologies, Eindhoven, Netherlands). The CV curves were obtained at a scan rate of 10 mV·s^− 1 and a potential step of 2 mV. The electrochemical measurements were performed in a three-electrode system comprising a glassy carbon working electrode, Ag/AgCl (saturated KCl solution) reference electrode, and a Pt wire counter electrode. BTP was coated on the working electrode using drop casting method for CV measurements. 1×PBS solution (measured pH = 7.24) was used as the supporting aqueous electrolyte. Before measurements, all electrolyte solutions were degassed with argon gas to prevent interference associated with dissolved O₂. We primarily measured oxidation and reduction potential (vs. Ag/AgCl) from on-set values, then converted the values to potential versus the normal hydrogen electrode (NHE) by adding + 0.197 V to obtain E^(0/−) and E^(+/0). The values are E^(0/−) (− 0.89 V vs. NHE) and E^(+/0) (1.00 V vs. NHE).

Excited redox potentials of BTP.

The photocatalytic redox potential was calculated using generally used method⁵¹. As described above, E^(0/−) (-0.89 V vs. NHE) and E^(+/0) (1.00 V vs. NHE) values were measured based on the CV results. To estimate the photocatalytic activity of BTP, E*^(0/−) (1.47 V vs. NHE) and E*^(+/0) (-1.36 V vs. NHE) values were also calculated by adding and subtracting the E^(0/0) value (2.36 eV), respectively.

Hydroxyl radical assay (hydroxyphenyl fluorescein, HPF).

Hydroxyl radicals were detected using an HPF assay. HPF (Invitrogen) is an indicator of hydroxyl radicals and peroxynitrite. The hydroxyphenyl group of HPF is eliminated by the hydroxyl radical, and HPF is subsequently converted to the fluorescent form (fluorescein). Thus, we prepared aqueous BTP (5 µM) and HPF solutions (5 µM) and measured fluorescence at 515 nm under various conditions: (1) before and (2) after BTP photoactivation in an Ar-bubbled aqueous solution, (3, 4) in a normoxic aqueous solution, (5, 6) in DMF solution, and (7, 8) in 23 mM H₂O₂ solution. In this assay, the final concentration of BTP was 5 µM. Furthermore, blue LED (λ_max = 450 nm) (HepatoChem Inc., USA) was used to photoactivate BTP (16.6 mW·cm^− 2 for 2 min = 2 J·cm^− 2), and all conditions except for the normoxic condition (3, 4) were maintained in conjugation with Ar bubbling conditions. A microplate reader (SpectraMax M5e, USA) was used to measure fluorescein fluorescence. Results were obtained using three distinct experimental samples.

Electron paramagnetic resonance (EPR) spectroscopy.

EPR spectroscopy with a spin trap was employed to identify the ROS generated by BTP photocatalysis because spin-adducts show various EPR spectra depending on the type of ROS. All EPR spectra were obtained in an aqueous solution at room temperature with 5-tert-butoxycarbonyl-5-methyl-1-pyrroline N-oxide (BMPO). The BTP stock solution (20 mM in DMF) and H₂O₂ were added to the BMPO aqueous solution to prepare 1 mM BTP, 10 mM H₂O₂, and 10 mM BMPO aqueous solution (H₂O:DMF = 95:5, v/v). The solution was irradiated with white room light for 5 minutes, and then transferred to a capillary EPR tube to measure the EPR spectra. EPR measurements were performed at Korea Basic Science Institute (KBSI) in Seoul, Korea. X-band (9.6 GHz) EPR spectra were derived using a Bruker EMX Plus 6/1 spectrometer equipped with a dual-mode cavity (ER 4116DM). The spectra were obtained using the following experimental parameters: microwave frequency, 9.6 GHz; microwave power, 2.9 mW; modulation amplitude, 1 G; time constant, 20.48 ms; 16 scans.

Hydrogen peroxide assay.

The hydrogen peroxide generation by BTP photoactivation was measured by the horseradish peroxidase (Sigma Aldrich, USA) and N,N-diethyl-p-phenylenediamine (DPD) (Sigma Aldrich, USA). First, the stock solution of peroxidase (1 mg·mL^− 1 in DI water) and DPD (50 mM in 1 M H₂SO₄ aqueous solution) were prepared. Then, the three 50 µM BTP solutions (in normoxic PBS, Ar-bubbled DMSO, and Ar-bubbled PBS) were irradiated by the blue LED (λ_max = 450 nm, 66.7 mW·cm^− 2) (HepatoChem Inc., USA). During the light exposure, assay samples were obtained at 30-minute intervals up to 150 minutes. The 200 µL of each sample solution was dissolved in the 224 µL DI water, and 80 µL of sodium phosphate buffer (pH 6) was added to each sample solution. Then, 10 µL of DPD and peroxidase stock solution was added. Right after that, the absorbance at 551 nm of each sample solution was measured. The results were obtained from three distinct experimental samples.

Cell culture.

HeLa, Panc-1, and MiaPaca-2 cells were purchased from ATCC and grown on the cell culture plates containing 90% Gibco™ DMEM (Thermo Fisher, USA), 10% fetal bovine serum (FBS), 50 units·mL^− 1 of penicillin, and 50 µg·mL^− 1 of streptomycin. The cells were grown at 37°C in a humidified atmosphere containing 5% CO₂. Wild-type and GSDMD-/- immortalised bone-marrow-derived macrophages (iBMDM) were obtained from the James Vince Lab of Walter and Eliza Hall Institute of Medical Research (WEHI). iBMDMs were grown on a cell culture plate with culture media comprising of 90% Gibco™ DMEM (Thermo Fisher, USA), 10% fetal calf serum (FCS), 50 units·mL^− 1 of penicillin, and 50 µg·mL^− 1 of streptomycin. The cells were grown at 37°C in humidified atmosphere containing 5% CO₂.

Membrane protein expression and purification.

The pTrcHisA vector containing the gene for E. coli GlpG membrane protein was transformed into BL21-Gold (DE3) pLysS (Agilent) ²⁶. A selected colony from the transformed agar plate was used to inoculate 10 ml of Luria-Bertani (LB) medium preculture with ampicillin (100 mg·mL^− 1) and grown overnight at 37°C. The preculture was added to the 1L LB medium containing 100 mg·mL^− 1 ampicillin and grown at 37°C. At an OD600 ≈ 0.7, the cell culture was induced by 0.4 mM Isopropyl β-D-thiogalactoside (IPTG) and further grown at 37°C for 3 hrs. Cells were harvested by centrifugation at 5700 rpm for 10 mins at 4°C. The cell pellet was resuspended in 25 mM Tris-HCl (pH 7.4),150 mM NaCl,1 mM TCEP, 10% Glycerol, 1 mM PMSF, and then lysed by using Emulsiflex C3 (Avestin) high-pressure homogenizer (~ 17000 psi). The cell lysate was mixed with n-dodecyl-β-D-maltoside (DDM) in the final concentration of 1% and kept in slow rotation for 1 hr at 4°C followed by centrifugation at 18000 rpm for 30 mins at 4°C. The supernatant was saved, and imidazole was added to the final concentration of 40 mM. For affinity column purification, Ni-IDA resin (Takara Bio) was washed with 1 ml with Tris-HCl (pH 7.4), and 150 mM NaCl was added and incubated for 1 hr. The binding solution was loaded to a gravity column and washed with 10 ml of 25 mM Tris-HCl (pH 7.4), 150 mM NaCl, 1 mM TCEP, 0.1% DDM, 10% Glycerol, and 40 mM imidazole three times. The protein sample was then eluted with 25 mM Tris-HCl (pH 7.4), 150 mM NaCl, 1 mM TCEP, 0.1% DDM, 10% Glycerol, and 300 mM imidazole. Eluted fractions were mixed, concentrated to ~ 500 µL with an Amicon 10K centrifugal filter device (Merck Millipore), and then further purified by size exclusion chromatography (Superdex 200 Increase 10/300 GL, Cytiva). Purified GlpG membrane protein in 25 mM Tris-HCl (pH 7.4), 150 mM NaCl, 1 mM TCEP, 0.1% DDM, and 10% Glycerol was stored at − 80°C.

Membrane protein stability assay.

4 µM GlpG was mixed with 100 ~ 150 µM BTP and the mixture was exposed to a blue light (λ_peak = 450 nm, 16.67 mW·cm^− 2) for 30 mins. Negative control samples without BTP and/or light were also prepared. 15% SDS-PAGE was used for the analysis, and the reduced gel band intensities for the destabilized protein by irreversible aggregation were quantified and normalised by the negative control with no BTP and no light. The results were obtained from three distinct experimental samples. For thermal denaturation, the sample mixture of GlpG and BTP exposed to the blue light for 30 mins was aliquoted in each 15 µL and incubated at various temperatures of 30.0, 45.0, 55.0, 65.0, 77.1, and 89.9°C for 10 mins, followed by cooling to 10°C in Blue-Ray Biotech Thermal Cycler. The thermal-shocked samples were centrifuged at 17,000 g at 4°C for 30 mins, and then the supernatants were analysed with 15% SDS-PAGE. The gel band intensities for the various temperature conditions were normalised by the one for a normal sample stored at 4°C. The results were obtained from three distinct experiment samples.

Single-molecule tweezer assay.

The single-molecule forced-unfolding assay was performed on a single-molecule magnetic tweezer apparatus that was custom-built previously described^34,35. Sample preparation of DNA-handled GlpG and a robust single-molecule system assembly for the force application was previously reported³⁶. The lipid bilayer environment for the membrane protein was reconstituted with a bicelle nanostructure, a lipid bilayer disc composed of DMPC lipid and CHAPSO detergent at a 2.5:1 molar ratio^30,35. Once singly tethered GlpG is found, repetitive force scanning from 1 pN to 50 pN and then back to 1 pN was applied by moving a pair of magnets toward and back from the sample chamber surface (0.3 mm·s^− 1). Extension change of the molecular construct as a response to force was measured by 3D-tracking the attached magnetic bead. Waiting for 120 secs at 1 pN between every pulling cycle was allowed for the refolding of unfolded GlpG. The negative control experiment was performed in 50 mM Tris (pH 7.5), 150 mM NaCl, and 2.0% bicelle under a blue light (λ_peak = 450 nm, 9.16 mW·cm^− 2). After three to four unfolding/refolding cycles, 10 ~ 20 µM BTP was injected into the sample chamber. A negative control experiment under an infrared light (λ_peak = 850 nm, 39.51 mW·cm^− 2) was also performed. The time durations (BTP injection ~ permanent damage on protein stability) and unfolding forces were collected and analysed statistically. The results were obtained from distinct experimental samples (n > 15).

LC-MS/MS.

Dry tryptic peptides were analysed using LC-MS/MS. A Q Exactive Plus Orbitrap mass spectrometer (Thermo Fisher Scientific, MA, USA) incorporated with a nanoelectrospray ion source was used for analyses. A C18 reverse-phase HPLC column (500 mm × 75 µm ID) was used to separate pure analyte from the crude peptide suspension. An acetonitrile/0.1% formic acid gradient of 2.4–24% was used as eluent at a flow rate of 300 nL/min. For MS/MS analysis, precursor ion scan MS spectra (m/z 400~–2000) were acquired with an internal lock mass. The 20 most intense ions were isolated via high-energy collision-induced dissociation. The mass spectrometry proteomics data were deposited to the ProteomeXchange Consortium via the PRIDE partner repository with the dataset identifiers PXD038746 and 10.6019/PXD038746.

LC-MS/MS data processing.

All MS/MS samples were analysed using Sequest Sorcerer platform (Sagen-N Research, San Jose, CA, USA). Sequest was set to search for Homo sapiens (20612 entries, UniProt (http://www.uniprot.org)), which includes frequently observed contaminants assuming the action of digestion enzyme trypsin. Sequest was searched with a fragment ion mass tolerance of 1.00 Da and parent ion tolerance of 10.0 PPM. The carbamidomethyl of cysteine was specified as a fixed modification in Sequest. Oxidation of methionine and acetyl at the N-terminus were specified as variable modifications in Sequest. Scaffold Q+ (version 5.1.0, Proteome Software Inc., Portland, OR) was used to validate MS/MS based peptide and protein identification. A peptide with a probability of higher than 99% for achieving an FDR of lower than 1.0% based on the no Scaffold Local FDR algorithm was accepted as true identification. A protein identification with a probability of higher than 14.0% for achieving an FDR of less than 1.0% and containing two or more identified peptides was accepted. Protein Prophet algorithm⁵² was used to calculate the protein probabilities. Proteins that contained similar peptides and could not be differentiated by MS/MS analysis alone were grouped to satisfy the principles of parsimony. The GO annotations for the proteins were retrieved from the NCBI database (downloaded on 11 February 2021). The total precursor intensity of the final proteomic data was used for label-free quantification to assess oxidation. Normalisation was performed iteratively on the intensities, while medians were used for averaging. Of the 5173389 spectra in the experiment at the given thresholds, 2120870 (41%) were included in the quantification. The values were log₂-transformed, pruned of those matched to multiple proteins, and non-reproducibly detected values were filled by imputed values representing a normal distribution around the detection limit. A new distribution was created by a Gaussian distribution with a downshift of 1.8 and width of 0.3 standard deviations. All processes were conducted using the Perseus software platform of Max Planck Institute of Biochemistry.

Assessment and visualisation of oxidised proteins.

The pre-determined significantly oxidised proteins were analysed using GOnet based on their log₂FC and log₂P⁵³. GO enrichment analysis with a q-value threshold of ≤ 0.05 yielded P values for GO terms. The terms of the top 20 non-zero FDR-adjusted -logP values were selected to distinguish biological processes that are affected by BTP photocatalysis. Oxidised proteome networks by functional association were determined using STRING and visualised using Gephi (ver. 0.9.2) and Cytoscape (ver. 3.9.1)⁵⁴. The crystal structures of oxidised proteins containing oxidatively modified Met residues are illustrated in PyMOL2 (2.5.2). Representative crystal structures were retrieved from reported literature in the Protein Data Bank (PDB). The PDB IDs of the structures demonstrated in this report were 3AHQ for 3RO1A, 3S7O for STT3A, 2OT3 for RAB21, and 1P49 for STS. Oxidized Met residues were assigned based on the LC-MSMS results.

LDH assay and ATP release assay.

The LDH-Glo™ Cytotoxicity Assay (Promega J2380, USA), LDH assay kit (ab65393, Abcam), and ATP determination kit (Invitrogen 2409086, USA) were used to detect the release of LDH and ATP. The samples for these assays were prepared in the same manner as that for the ELISA experiment. To set the maximum LDH release control, HeLa cells were lysed with 0.2% Triton X-100 for 15 min, and the medium was used for 100% LDH release. For the LDH assay, HeLa cells were grown with Liproxstatin-1 (10 µM) (Sigma Aldrich SML1414, USA) and z-VAD-fmk (4 µM) (Promega G7231, USA) for 16 h. BTP was then added to the cells (8 µM), followed by incubation for 2 h. After replacing the growth medium with a serum-free medium, the cells treated with the BTP/inhibitor were exposed to blue LED light (λ_max = 450 nm, 3 J∙cm^− 2). Two hours after light exposure, samples were obtained from the media for the LDH assay. For the LDH assay with GSDMD^−/− iBMDM, WT and GSDMD^−/− iBMDM were grown on cell culture dishes for one day. iBMDMs were incubated with BTP (8 µM) for 2 h, and then the cells were exposed to blue LED light (λ_max = 450 nm, 10 J∙cm^− 2) after the media was changed. Two hours after irradiation, samples were obtained from the media for the LDH assay. Results were obtained using three distinct experimental samples.

Western blot.

To investigate the activation of GSDMD and caspases by BTP photoactivation, we conducted western blot analysis with the following antibodies: GSDMD antibody, NBP2-33422 (Novus Biologicals); caspase-1 antibodies, ab207802 (Abcam) and PA5-29342 (Invitrogen); caspase-3 antibody, ab32351 (Abcam); caspase-4 p20 antibody, A94799 (Antibodies); caspase-5 antibody, sc-393346 (Santa Cruz); IL-1β antibody, P420B (Invitrogen); and IL-18 antibody, PA5-79479 (Invitrogen). HeLa cells incubated with BTP (10 µM) for 2 h were irradiated with a blue LED (λ_max = 450 nm, 3 or 10 J·cm^− 2), and the cells were further incubated for 2 h in a serum-free media. After incubation, the media were obtained and concentrated using an Amicon® Ultra-4 Centrifugal Filter, and a HaltTM protease inhibitor cocktail was added. The remaining cells were lysed with RIPA protein extraction solution (50 mM Tris-HCl pH 7.5, 150 mM NaCl, 1% NP-40, 0.5% deoxycholic acid, 0.1% SDS, and 1 mM PMSF) containing HaltTM protease inhibitor cocktail. The lysates and media were stored at -80 ℃ until further use. For western blotting, the protein concentration of each sample was determined using a BCA Protein Assay (Thermo Fisher 23227, USA). After protein denaturation with SDS-PAGE loading buffer (Biosesang, Republic of Korea) for 7 min at 70°C, the loaded protein solutions were separated by SDS-PAGE gel electrophoresis, and the separated protein bands were transferred onto a 0.2 µm polyvinylidene difluoride (PVDF) blotting membrane (GE Healthcare, Germany). Proteins on the membrane were blocked with 4% skim milk in 0.1% Tween-20 in Tris-buffered saline (TBST) for an hour. Subsequently, the membrane was incubated with the primary antibodies at a ratio of 1000:1 (500:1 for caspase-5 antibody) in 4% skim milk for 16 h at 4 ℃. After washing thrice with TBST (for 10 min each), the membrane was further incubated with the appropriate secondary antibody (3000:1 dilution in TBST), which are listed as follows: anti-mouse HRP (Invitrogen 31430, USA) or anti-rabbit HRP (Abcam ab205718, USA) After washing thrice, western blot chemiluminescence was measured using a ChemiDoc™ MP imaging system (Bio-rad, CA, USA) after development with Clarity reagent (Bio-rad, CA, USA). After the experiment, the antibodies attached to the membrane were eliminated using Restore Western Blot Stripping Buffer (Thermo Fisher, 21059). Western blotting was repeated using an antibody against β-actin (Invitrogen MA5-15739, USA). The experiment was repeated using negative controls (without BTP and/or light exposure).

Data availability

The authors declare that the data supporting the findings of this study are available within the article and its Supplementary Information. The protein information utilized in this study is available from the Homo sapiens protein sequence database (20612 entries, UniProt (http://www.uniprot.org)). Raw mass spectrometry dataset used for oxidised proteome analysis have been deposited to the ProteomeXchange Consortium via the PRIDE partner repository with the dataset identifier PXD038746 and 10.6019/PXD038746. Extra data are available from the corresponding author upon reasonable request. The source data underlying Fig. 1c−f; Fig. 2b, d, f−h; Fig. 3a−c, e, f; Fig. 4c; Fig. 5b, c, e−h; Extended Data Fig. 1a, b, e−j; Extended Data Fig 2e, g; Extended Data Fig. 4b; Extended Data Fig. 5; Extended Data Fig. 6; Extended Data Fig. 7b, c, and e are provided as a Source Data file.

Acknowledgements

This work was supported and funded by the National Research Foundation of Korea (NRF- 2021R1A2C2009504, 2021M3H4A1A03051390, 2021R1F1A1047853, and NRF-2020R1C1C1003937), National Cancer Center (NCC) (Research Fund HA22C010100), Korea Technology & Information Promotion Agency for SMEs (TIPA) (grant S3198656), New Renewable Energy Core Technology Development Project of the Korea Institute of Energy Technology Evaluation and Planning (KETEP) granted financial resource from the Ministry of Trade, Industry & Energy, Republic of Korea (No. 20223030010240), Korea Electric Power Corporation (grant R21XO02-8), and Korea Innovation Foundation (2021-IT-RD-0338). This work also was supported by the Carbon Neutral Institute Research Fund (1.220098.01) of UNIST and Research Fund (1.220033.01 and 1.190147.01) of UNIST. C. G. L. acknowledges support from the Global Ph.D. Fellowship Program through the National Research Foundation of Korea (NRF) funded by the Ministry of Education (NRF-2018H1A2A1061237). E. H. acknowledges support from the Basic Science Research Program through the NRF funded by the Ministry of Education (2.220901.01). J. E. V. was funded by a National Health and Medical Research Council (NHMRC) of Australia ideas grant (1183070) and investigator grant (1172929).

Author contribution

C. L. and T.-H. K. conceived and conceptualised this study. C. L., M. P. and T.-H. K wrote the manuscript. C. L., M. P., and B. W. contributed equally to this work. C. L. contributed to all aspects of this study in conception, conduction, and analysis of experiments, in particular organic synthesis, spectroscopic analysis, photocatalysis design, ROS assays, confocal and SIM imaging, cell viability assay, and immunoblots. M. P. performed sampling and analysis of oxidised proteome and participated in cytotoxicity tests, hypoxia application, and immunoblots. J. K. S. reviewed and supported all proteomics. B. W. and D. M. conceived and conducted protein stability assays and single-molecule tweezer experiments. C. G. L. performed MTT assays and flowcytometry. E. H. conceived and performed electrochemical analysis for amino acid oxidation. G. Y. and J. K. L. performed organic synthesis and characterisation. C. L. and G. Y. performed ROS assay. J. K. L. conducted TSCPC analysis. D.-H. R. performed cyclic voltammetry. Y. H. K performed MTT assay under hypoxia. J. Y. participated in immunoblots and cell experiments. J. E. V. and S. A. H generated and provided immortalised WT and GSDMD knock out iBMDMs and reviewed the manuscript. T.-H. K. supervised all aspects of this study.

Competing interests

The authors declare no competing interest.

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Oxidative photocatalysis on membranes triggers non-canonical pyroptosis

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