Natural photosynthetic system for restoring homeostasis of animal organelle interaction network

Xia, Chen; Dai, Zhanqiu; Wang, Yongcheng; Yu, Jiachen; Wang, Zijie; Huang, Zhenxiang; Pan, Kaifeng; Yang, Ke; Chen, Yiyu; Zhong, Peiyu; Gu, Chenhui; Fan, Shunwu; Lin, Xianfeng; Chen, Pengfei

doi:10.1038/s41467-026-69825-y

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Published: 31 March 2026

Natural photosynthetic system for restoring homeostasis of animal organelle interaction network

Chen Xia ORCID: orcid.org/0000-0001-8901-0827^1,2,3^na1,
Zhanqiu Dai ORCID: orcid.org/0000-0002-7408-7909^1,3,4^na1,
Yongcheng Wang^1,3^na1,
Jiachen Yu^1,3,
Zijie Wang^1,3,
Zhenxiang Huang^1,3,
Kaifeng Pan^1,3,
Ke Yang^1,3,
Yiyu Chen^1,3,
Peiyu Zhong^1,3,
Chenhui Gu ORCID: orcid.org/0000-0002-8698-9395^1,3,
Shunwu Fan ORCID: orcid.org/0000-0003-3443-4395^1,3,
Xianfeng Lin ORCID: orcid.org/0000-0003-1871-774X^1,3,5 &
…
Pengfei Chen ORCID: orcid.org/0000-0003-1484-2850^1,3,5

Nature Communications volume 17, Article number: 3087 (2026) Cite this article

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Abstract

Disruption of organelle interactions due to metabolic stress is a crucial factor in the pathological processes of many degenerative diseases. Compared with animal cells, the participation of chloroplasts enables plant cells to show defensive adaptation under stress. Therefore, delivering plant-derived photosynthetic systems into animal cells may help to establish a more stable organelle interaction network. Here, we show that plant-derived photosynthetic systems can effectively restore homeostasis of the interaction network of animal organelles. Specifically, plant-derived nanothylakoid units provide energy to animal cells, regulate intracellular Ca²⁺ homeostasis, increase endoplasmic reticulum (ER) lipid unsaturation and global membrane fluidity, reduce abnormal contact between mitochondria and ER, and alleviate mitochondrial dysfunction. By combining implantable light-emitting diodes with wireless charging, we expand photosynthesis therapy, enabling treatments for deeper tissues. This study provides a proof-of-concept for disease treatment based on the regulation of organelle interaction networks by natural photosynthetic systems and establishes a therapeutic approach for treating deep tissues.

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Introduction

The early evolution of life underwent a process of formation from simple organic molecules to precellular and primitive lives^1,2. For the evolution of complex life, the acquisition of mitochondria in eukaryotes is a prerequisite^3,4. The endosymbiotic event that gave rise to mitochondria enabled a substantial expansion in gene expression and protein synthesis in eukaryotes relative to prokaryotes. This increase was driven by the emergence of specialized bioenergetic membranes and enhanced ATP-producing capacity, which together provided the energetic foundation for the biological diversity and complexity^4,5. However, different types of diseases take a huge toll on life by placing stress on cells. Plants possess mitochondria and chloroplasts and exhibit unique defensive adaptations to stress^6,7. Malate/oxaloacetate translocators and malate dehydrogenases form the malate valve, which metabolically links chloroplasts and mitochondria by transferring excess reducing equivalents from the chloroplast to maintain stromal redox balance. Under stresses such as high light, elevated CO₂, cold, drought, or salinity, malate valve activity adjusts to maintain ATP/NAD(P)H homeostasis, limit ROS formation, and support stress resilience^8,9,10. Additionally, plants also enhance the unsaturation of fatty acids in membrane lipids that preserves membrane fluidity and sustains the activity of essential membrane-associated proteins under cold stress^{11,12,13,14,15,16,17,18}. Together, these redox-balancing and membrane-stabilizing mechanisms enable plants to maintain metabolic function and tolerate abiotic stress. However, compared with plant cells, animal cells lack the ability to tolerate different stressful environments. Therefore, transplanting plant-derived photosynthetic systems into animal cells may enhance their stress tolerance.

A previous study has used cell membrane camouflage encapsulation to deliver plant-derived nanothylakoid units (NTUs) into animal cells to control the deficient anabolism of pathological animal cells under light¹⁹. Healthy cells usually have a highly sophisticated spatiotemporal architecture, enabling precise organization of dense organelles and macromolecular complexes, accurate spatial intersections, and timely metabolite exchange²⁰. However, as miniature organelles that restore the functions of pathological animal cells, whether NTUs can transplant the defensive adaptation capabilities of plants into animal cells remains unclear.

The premise of organelle interactions is the proximity between the organelles²¹. As the largest organelle in eukaryotic cells, the endoplasmic reticulum (ER) forms a network throughout the cell, maintains many membrane contact sites, and acts as a hub for intracellular protein and lipid synthesis²². The ER dedicates a special domain for metabolic interaction with mitochondria and is called the mitochondria-associated ER membrane (MAM)²³. MAMs play important roles in various cellular functions, including acting as a tight platform for lipid synthesis and lipid transfer between the ER and mitochondria^23,24. Mitochondria, the energy factories of eukaryotic cells, support the cascade of organelle interactions by generating energy²⁵. The contact sites between the outer mitochondrial membrane and ER (MERCS) ensures spatiotemporal communication between mitochondria and the ER^26,27. Animal cells exhibit ER–mitochondria contact alterations under stress^28,29. Targeting individual organelles may not systematically correct dysregulated organelle interactions, and it is urgent to identify methods to restore these dysregulated interaction networks. Intervertebral disc degeneration (IVDD) is a degenerative disease of the deep retroperitoneal tissues that disrupts ER and mitochondrial functions in nucleus pulposus cells (NPCs), which are crucial pathological factors in its pathogenesis^30,31. Given the central position of the ER-mitochondrial network in cell metabolism and because NTUs participate in cellular metabolism by producing energy and reducing equivalents¹⁹, NTUs may have the potential to protect dysregulated organelle interaction networks under pathological conditions.

Selecting the appropriate light to initiate the photosynthetic reaction is essential to achieve the in vivo therapeutic applications of NTUs. However, commonly used external light sources (lasers, light-emitting diodes, and lamps) are limited by poor penetration of light into tissues. Previous studies have attempted to introduce optical fibers into the body through endoscopy to achieve light transmission^32,33; however, the wires required for optical fibers are rigid, fragile, bulky, and difficult to stabilize. Wireless power transfer involves transmitting electrical energy using a cable-free method, and this method is used to deliver controlled doses of light into deep tissues for therapeutic purposes^34,35.

In this work, we aim to explore whether plant-derived NTUs can restore animal cell homeostasis under pathological conditions by modulating the ER–mitochondria interaction network. The IVDD animal model is used for conceptual verification. Similarly, we use an NPC membrane to nanoencapsulate NTUs (NM-coated NTUs, NM-NTUs), achieving light-controlled regulation of the organelle interaction network in rat IVDD models. Moreover, to simulate deep-tissue treatment in clinical practice, we design a wirelessly powered light-emitting diode (wp-LED) as an implantable light source and combine it with a wireless controller connected to a smartphone to remotely control the photosynthetic response of NTUs in deep tissues. The efficacy of combining wp-LED with NTU injection is validated in rabbit IVDD models (Fig. 1). In summary, we find that implanting plant-derived photosynthetic systems into mammalian cells can effectively restore homeostasis of the ER–mitochondria interaction network. By combining implantable wp-LED, we enable more effective treatments for deeper tissues.

**Fig. 1: Schematic illustration of this study.**

Results and discussion

Disruption of the ER-mitochondria interaction network in human IVDD

ER stress and mitochondrial metabolic imbalance have been proposed to be involved in the pathological process of IVDD^36,37. Transcriptomic analyses were performed on normal and degenerative human nucleus pulposus (NP) (Fig. 2a). The Pfirrmann degeneration grade indicates the degree of disc degeneration between the two groups (Supplementary Fig. 1a). Principal component analysis (PCA) showed that the genes in the normal group were distinct from those in the degeneration group (Fig. 2b). Compared with the normal group, 799 genes were differentially expressed in the degeneration group, with 369 upregulated and 430 downregulated (Fig. 2c). Gene expression patterns in the degeneration group showed a significant increase in the expression of oxidative stress-related (CYBB, RAC2, and EDN1) and ER stress-related (IGFBP1, STC2, and PTX3) genes (Fig. 2d). To further determine the levels of ER- and mitochondrial-related pathway enrichment, we performed gene set enrichment analysis (GSEA). The degeneration group exhibited an upregulated expression of genes involved in the reactive oxygen species (ROS) and reactive nitrogen species (RNS) production, ER stress, calcium import into the mitochondrion, and mitochondria permeability (Fig. 2e; Supplementary Fig. 1b, c).

**Fig. 2: Disruption of the endoplasmic reticulum (ER)-mitochondria interaction network in human intervertebral disc degeneration (IVDD).**

To validate the findings of transcriptomic analyses and study cellular changes of human NPCs, we established a primary culture of normal and degenerated human NPCs. On the basis of previous reports, the phenotypic gene expression of human NPCs changes significantly after approximately 11–13 days of in vitro culture^38,39. To evaluate the impact of in vitro culture on the changes in the phenotype of degenerated NPCs in this study, we monitored the phenotypic profiles of degenerated human NPCs cultured for 1–9 days. Referring to previous studies^38,39, we examined the following NPC phenotypic markers: genes (CD24, KRT18, KRT19, SOX9, MIA, FN1, COL1A1, COL2A1, and COL3A1) and proteins (CD24 and KRT19). The results indicated that the phenotype of degenerated NPCs remained stable during the first 5 days of culture, whereas changes in the expression of certain markers began to emerge on day 7 (Supplementary Fig. 2a, b). Therefore, a 5-day culture time was selected in this study to reduce the impact of phenotypic changes on subsequent experiments. Next, we measured cell viability to evaluate the cellular status. In the 3-(4,5-dimethylthiazol-2-yl)-2,5-diphenyltetrazolium bromide (MTT) assay, the cell viability of degenerated human NPCs decreased to 68.2% compared with that of normal human NPCs (Supplementary Fig. 3a). However, in the live/dead staining assay, we detected no significant difference in the percentage of live cells between the degenerated NPCs and normal NPCs (Supplementary Fig. 3b, c). These findings suggested that although degenerated NPCs exhibited reduced metabolic activity and viability, substantial cell death did not occur. These results also suggested that the in vitro model matched IVDD, where a large percentage of NPCs were alive and that the findings in the following series of experiments were not due to a change in cell number but rather cellular metabolic changes.

The ER, the primary storage organelle for intracellular Ca²⁺, pumps cytoplasmic Ca²⁺ through sarcoplasmic/ER Ca²⁺-ATPase. Fluo-4/AM (intracellular Ca²⁺ indicator) and Rhod-2/AM (mitochondrial Ca²⁺ indicator) staining results showed that cytosolic Ca²⁺([Ca²⁺]_c) and mitochondrial Ca²⁺([Ca²⁺]_m) levels were significantly increased in degenerative NPCs (Fig. 2f). Since the ER is the primary intracellular Ca²⁺ store, the rise in [Ca²⁺]_c and [Ca²⁺]_m may reflect a decrease in the Ca²⁺ concentration within the ER ([Ca²⁺]_ER). Ca²⁺, an ER signaling molecule, participates in protein folding inside the ER. A decrease in the [Ca²⁺]_ER may cause ER stress through an unfolded protein response⁴⁰. Our results showed that degenerative NPCs had increased levels of ER stress-related proteins (CHOP and ATF4) (Fig. 2g). ER stress causes increased MERCS and increased mitochondrial Ca²⁺ uptake from the ER⁴¹. To further clarify the ER–mitochondrial interaction network, we assessed the MERCS structure of normal and degenerative NPCs by immunostaining the ER and mitochondria with calnexin and translocase of the outer mitochondrial membrane 20 (TOMM20), respectively. The results revealed increased colocalization of the ER and mitochondria in degenerative NPCs (Fig. 2h, i). This was confirmed using transmission electron microscopy (TEM). ER–mitochondria contacts can be characterized by gap distances, which are usually in the range from 10–80 nm^42,43. In accordance with a previous study⁴⁴, we defined contact sites as those with tethered proximity between membrane-bound organelles. The contacts were categorized and quantified into three distinct ranges: ≤25 nm, 25<distance≤50 nm, and 50<distance≤100 nm. Compared with normal NPCs, degenerated NPCs presented significantly increased ER tubules closely apposed to mitochondria, indicating increased formation of MERCS (Fig. 2j, k). There was no significant difference in the number of mitochondria or the average length of ER tubules between normal and degenerated NPCs (Fig. 2l, m). The MERCS is crucial in regulating Ca²⁺ homeostasis from the ER to the mitochondria. Aberrant MERCS formation promotes [Ca²⁺]_m overload^45,46 and triggers the permeability of the inner mitochondrial membrane by opening the mitochondrial permeability transition pore (mPTP)⁴⁷. Direct assessment of mPTP opening through the calcein release⁴⁸ assay confirmed increased mPTP opening in degenerative NPCs (Fig. 2n). mPTP opening caused the depolarization of the mitochondrial membrane potential and oxidative stress⁴⁹. Degenerated NPCs showed higher ROS production (Fig. 2o) and lower mitochondrial membrane potential than normal NPCs (Fig. 2p). Taken together, the disrupted ER-mitochondrial interaction network plays a crucial role in the process of IVDD, and restoring the dysfunctional organelle interaction network may ameliorate IVDD.

Preparation and characterization of NM-NTUs

The above results reveal that under pathological conditions, the ER-mitochondrial interaction network is disrupted. Given that plants possess chloroplasts and mitochondria and show defensive adaptations to stressful environments^6,7, we used our previously constructed cell membrane-encapsulated NTUs, which can be transplanted across species¹⁹, to investigate their potential to regulate the interaction network of animal organelles. We coextruded the purified NPC membranes and NTUs to obtain NM-NTUs. To evaluate the encapsulation efficiency of the NTUs by NM, we used nanoscale flow cytometry (nanoFCM). The nanoFCM data indicated that the encapsulation efficiency was dependent on the initial ratio of NTUs to NMs. At the optimal formulation ratio (NTU to NM protein ratio of 1:150), 77.83 ± 4.01% of the NTUs could be encapsulated into NMs (Supplementary Fig. 4a), with approximately 51.37 ± 2.08% of the NMs successfully loaded with NTUs (Supplementary Fig. 4b). This ratio was used in subsequent experiments. The NM-NTU particle size was approximately 229 nm (Fig. 3a). The surface zeta potential results showed that the NTUs were coated with the cell membrane (Fig. 3b). The result of sodium dodecyl sulfate-polyacrylamide gel electrophoresis (SDS-PAGE) further confirmed the retention of the photosynthetic system proteins in NM-NTUs (Fig. 3c). The NM-NTU core-shell structure was observed using cryogenic transmission electron microscopy (cryo-TEM) (Fig. 3d). The cell counting kit-8 (CCK-8) assay revealed that the NM-NTU treatment had no evident cell toxicity (Fig. 3e).

**Fig. 3: Preparation and characterization of NPC-membrane-coated NTUs (NM-NTUs).**

Cell membrane-coated nanoparticles have unique characteristics such as immune escape, molecular recognition, and cell targeting^50,51. We evaluated the selective targeting ability of the NM-NTUs. NTUs were first labeled with DiI. In the intervertebral disc environment, the primary cells present are NPCs, annulus fibrosus cells (AFCs), and endplate chondrocytes (EPCs). Therefore, rat NPCs, AFCs, and EPCs were treated with NM-NTUs (NTUs labeled with DiI). NM-NTUs were more highly taken up by NPCs than by AFCs or EPCs (Fig. 3f, g), indicating that NM-NTUs were preferentially taken up by NPCs. To confirm that membrane fusion had occurred, rat NPCs^DiI (NPC membrane labeled with DiI) were treated with NM^DiO-NTUs (NM labeled with DiO; NTU unlabeled). Owing to the characteristics of cell membrane fusion, the fluorescence signal of NM^DiO colocalized with the fluorescence signal of NPC^DiI, confirming the occurrence of membrane fusion (Fig. 3h). Lysosomes are organelles that degrade macromolecules via various membrane-trafficking pathways⁵². Many pathogens avoid lysosomal degradation by escaping the endocytic pathway⁵². In our study, rat NPCs were coincubated with either NTU^DiI (NTU labeled with DiI) or NM-NTU^DiI (NM unlabeled; NTU labeled with DiI). The results revealed that in NPCs treated with NTU alone, the NTU^DiI fluorescence signal colocalized with that of the lysosomes (labeled with LysoTracker Green). In contrast, in NPCs treated with NM-NTU^DiI, only a few signals of NTU^DiI colocalized with lysosomes (Fig. 3i, j). These results indicated that the NM coating enabled the NTUs to bypass lysosomal degradation. Furthermore, the NTU’s post-internalization behavior after NM-NTU uptake was highly significant; the colocalization experiments revealed that the NTU signals colocalized with actin and tubulin to a high degree, and only a few NTU signals colocalized with the ER, mitochondria, and Golgi apparatus (Supplementary Fig. 5a, b). These results indicated that the NTU might achieve motility after entering the cell through the cytoskeleton. This phenomenon has similarities to previous reports of cytoskeleton-driven chloroplast movement in plants^53,54. Moreover, to visualize MERCS and NTU colocalization in NPCs, we transfected immortalized human primary NPCs (IH-NPCs) with a previously validated split-GFP-based ER–mitochondria contact site sensor (SPLICS)^55,56. Following the transfection of IH-NPCs with SPLICS, we treated them with NM-NTUs and performed confocal microscopy observations. Analysis of the colocalization between the MERCS signal (reconstituted GFP, green) and the NM-NTU signal (DiI labeled NTUs, red) revealed that the Mander’s coefficient for the percentage of NTUs that colocalized with MERCS was 5.29 ± 1.55% (Supplementary Fig. 6a–c). Taken together, the above results indicated that in the intervertebral disc microenvironment, NM-NTUs were preferentially taken up by NPCs. After entering NPCs, NTUs could evade lysosomal digestion and probably did not physically interact with MERCS.

NM-NTUs enhance NPC anabolism

Similar to that in other degenerative diseases, degenerated NPCs show an imbalance between anabolic and catabolic processes⁵⁷. The ATP generation ability of NTUs has been confirmed in our previous study¹⁹. Studies have used interleukin (IL)-1β to induce NPC degeneration, which may be related to its ability to increase the expression of matrix degradation-related genes (MMP3, MMP13, and ADAMTS-4) and decrease the expression of matrix synthesis-related genes (ACAN and COL2A1), resulting in an imbalance between catabolic and anabolic events in NPCs^58,59,60. Here, we measure the ability of NM-NTU to increase ATP levels in IL-1β-induced degenerated NPCs. We treated IL-1β-stimulated NPCs with a range of NM-NTU concentrations (0.125, 0.25, 0.5, 1.0, and 2.0 μg mL⁻¹ for chlorophyll). The results revealed that as the concentration of NM-NTU increased from 0.125 to 0.5 μg mL⁻¹ for chlorophyll, its ability to increase ATP in degenerative NPCs gradually increased, demonstrating the dose-dependent effects of NM-NTUs (Supplementary Fig. 7). A plateau was observed at NM-NTU concentrations higher than 0.5 μg mL⁻¹ for chlorophyll. On the basis of these results, we selected a concentration of 0.5 μg mL⁻¹ for chlorophyll in the subsequent in vitro experiments. Our further experiments also suggested that, compared with IL-1β group, the IL-1β + NM-NTU + dark group could not increase the intracellular ATP level (Fig. 4a). These results indicated that nonfunctional NM-NTUs (without light illumination) did not change the bioenergetics of cells and that light exposure was necessary for NM-NTUs to generate ATP in mammalian cells. To specifically quantify the ATP contribution from the NM-NTUs, we employed a pharmacological inhibition strategy to suppress endogenous cellular ATP production. We used 2-DG and rotenone to inhibit ATP production through glycolysis and oxidative phosphorylation in nucleus pulposus cells (NPC_ΔATP)^61,62. Moreover, we used NM-NTUs + light to increase the ATP content in NPC_ΔATP, hypothesizing that the increased ATP in these cells was derived mainly from NM-NTUs. The results revealed that the rate of ATP production by NM-NTUs in NPC_ΔATP was 8.50 ± 1.42 pmol min⁻¹ per 10⁵ cells (Supplementary Fig. 8). Moreover, results of quantitative real-time polymerase chain reaction revealed that NM-NTUs under light illumination significantly upregulated Col2a1, Acan, and Sox9 and decreased Mmp3, Mmp13, and Adamts5 mRNA levels in degenerated NPCs (Fig. 4b). Immunofluorescence showed results consistent with the upregulation of Col II and aggrecan and the downregulation of MMP13 and ADAMTS-5 protein levels in NPCs treated with NM-NTUs + light, indicating that NTUs could improve the anabolism of degenerated NPCs (Fig. 4c).

Persistent ATP deficiency in cells is crucial in the dysregulation of cell metabolism. We used RNA-Seq to determine the possible mechanisms and molecular pathways underlying the protective effects of NM-NTUs on NPCs. Compared with the control group, the IL-1β group and the IL-1β + NM-NTU + dark group presented a greater number of differentially expressed genes, whereas compared with the control group, the IL-1β + NM-NTU + light group presented a lower number of differentially expressed genes (Fig. 4d; Supplementary Fig. 9a–d). In addition, we compared differentially expressed genes from the transcriptomes of normal and degenerative human NPCs, as well as from NPCs with and without IL-1β treatment. The results suggested that IL-1β treatment upregulated the Adamts7 (matrix degradation-related gene⁶³) and Il1b genes and downregulated the Col2a1 (matrix synthesis-related gene) and Sesn3 (protection against oxidative stress⁶⁴) genes. These differentially expressed genes were identical in the human transcriptome and exhibited consistent trends in the degenerative group, strengthening the validity of the IL-1β-induced NPC degeneration model. We further evaluated the effects of NM-NTUs on degenerated NPCs. PCA revealed distinct clustering among samples within groups, with similar gene expression profiles in the control and IL-1β + NM-NTU + light groups, while both groups clustered separately from the IL-1β and IL-1β + NM-NTU + dark groups (Fig. 4e). Furthermore, in contrast to the IL-1β + NM-NTU + dark group, the IL-1β + NM-NTU + light group presented lower levels of ROS-related gene (Prdx6) and mitochondrial apoptosis-related genes (Nptx1 and Bid) (Fig. 4f). GSEA results indicated that, compared to the control group, the IL-1β group and the IL-1β + NM-NTU + dark group presented upregulated expression of genes related to ROS signaling, negative regulation of [Ca²⁺]_ER, and mitochondrial apoptosis and downregulated expression of genes related to ECM assembly (Supplementary Fig. 9e, f). In contrast to the IL-1β group and the IL-1β + NM-NTU + dark group, the IL-1β + NM-NTU + light group exhibited downregulation of genes associated with ROS signaling, negative regulation of [Ca²⁺]_ER, and mitochondrial apoptosis and upregulation of genes associated with ECM assembly (Fig. 4g; Supplementary Fig. 9g). Taken together, these results demonstrated that NM-NTUs could restore metabolic homeostasis in degenerated NPCs in a light-dependent manner and might have positive effects on the ER and mitochondria within NPCs.

NM-NTUs correct dysfunctional organelle interaction network

We further evaluated the effect of NM-NTUs on the interaction network between the ER and mitochondria. The ER is the primary store of cytosolic Ca²⁺, and the ATP produced by NTUs may enhance the ER Ca²⁺ storage function through energy supplementation. To validate this finding, we utilized the genetically encoded ER Ca²⁺ indicator (G-CEPIA1er)⁶⁵ to specifically monitor the ER intraluminal Ca²⁺ concentration ([Ca²⁺]_ER) and used 3-(3,4-dichlorophenyl)-1,1-dimethylurea (DCMU)⁶⁶ to disrupt light-dependent ATP synthesis by NM-NTUs. The results showed that the NM-NTUs + light treatment significantly increased [Ca²⁺]_ER compared with the NM-NTUs + dark treatment in 30 min (Supplementary Fig. 10a, b). Moreover, the inhibition of ATP production in NM-NTUs + light by the DCMU significantly attenuated the NM-NTU-induced increase in [Ca²⁺]_ER (Supplementary Fig. 10a, b). Next, we evaluated [Ca²⁺]_c using Fluo-4/AM dye. The NM-NTUs + light treatment decreased [Ca²⁺]_c compared with that in the NM-NTUs + dark group (Fig. 5a, b). In summary, these results suggested that NM-NTUs might restore intracellular and ER Ca²⁺ homeostasis. Because ER Ca²⁺ perturbation may cause ER stress, we examined the effect of NM-NTUs on ER stress and found that NM-NTU effectively reduced the abundance of ER stress-related proteins (IRE1, ATF6, ATF4, and CHOP) in degenerated cells (Fig. 5c). We subsequently assessed the formation of MERCS by immunostaining the ER and mitochondria and observed that the NM-NTU + light treatment resulted in a decrease in MERCS (Fig. 5d, e). We further evaluated the effect of NM-NTUs on the ER stress-related MERCS structure by TEM. Compared with the IL-1β or IL-1β + NM-NTU + dark group, the IL-1β + NM-NTU + light group had significantly fewer ER closely apposed to mitochondria (Fig. 5f, g). ER length and mitochondrial counts were calculated for each field, and the results revealed no significant difference in the number of mitochondria or the average length of ER tubules among the four groups (Fig. 5h, i). An aberrant increase in MERCS causes mitochondrial Ca²⁺ overload and impairs cellular metabolic homeostasis⁶⁷. As Ca²⁺ influx in the mitochondria can trigger mPTP opening, we measured [Ca²⁺]_m and found that NM-NTU + light treatment caused an evident decrease in [Ca²⁺]_m compared with that in the IL-1β group (Fig. 5j, k). Moreover, NM-NTU + light treatment significantly decreased the mPTP opening in degenerated NPCs (Fig. 5j, l). The continued opening of the mPTP may cause dissipation of the mitochondrial membrane potential and bioenergetic failure of the cell⁶⁸. Next, we examined the effect of NM-NTUs on the restoration of mitochondrial function and found that NM-NTU + light treatment increased the mitochondrial membrane potential in degenerated NPCs (Fig. 5m, n). To investigate whether NM-NTU treatment led to increased ATP production from endogenous mitochondria in degenerated NPCs, we transfected IH-NPCs with a genetically encoded mitochondrial ATP indicator (mito-AT1.03) to measure changes in mitochondrial ATP concentrations^69,70,71. The ratio of yellow fluorescence protein (YFP) to cyan fluorescence protein (CFP) was measured by flow cytometry to determine the mitochondrial ATP level. The results showed that following NM-NTUs + light treatment, the mitochondria within degenerated cells produced more ATP compared with the IL-1β group (Supplementary Fig. 11a, b). These results indicated that NM-NTUs might stimulate endogenous mitochondria in degenerated NPCs to produce ATP. In conclusion, while the NTUs themselves produced a quantifiable amount of ATP, a key therapeutic mechanism involved the NTU-generated ATP acting as a signal to restore the homeostasis of organelle interaction networks and promote the energy production of endogenous mitochondria. NTUs may also contribute to cellular metabolism through the generation of NADPH, which plays a critical role in redox homeostasis^19,72. We further evaluated the effects of NM-NTUs + light on the regulation of intracellular NADPH levels. The results revealed that compared with that in the control NPCs, the NADPH content in the IL-1β-induced degenerated NPCs was decreased (Supplementary Fig. 12a). The IL-1β + NM-NTUs + light treatment significantly restored cellular NADPH levels compared with the IL-1β + NM-NTUs + dark group (Supplementary Fig. 12a). Furthermore, we found that IL-1β + NM-NTUs + light treatment resulted in reduced intracellular ROS levels compared with the IL-1β + NM-NTUs + dark group (Supplementary Fig. 12b, c). These findings supported the conclusion that, in addition to producing energy, NTUs might also help to mitigate cellular oxidative stress through the generation of NADPH. Studies have shown that the ER-mitochondrial interface contains a nanodomain of H₂O₂ originating from mitochondrial cristae⁷³ and that high mitochondrial ROS levels promote MAM formation⁷⁴. We hypothesized that NADPH produced by NTUs might indirectly affect the contact between mitochondria and the ER through the regulation of redox homeostasis. Our results indicate that NM-NTUs can restore intracellular Ca²⁺ homeostasis, inhibit abnormal MERCS formation, and reestablish a stable ER-mitochondrial interaction network (Fig. 5o).

NM-NTUs remodel ER lipid composition

ER-localized enzymes synthesize most cellular lipids, and the ER has a major influence on cellular lipid biomass and membrane biogenesis⁷⁵. ER membrane lipid composition is altered in response to stressful conditions. For example, under prolonged nutrient and oxidative stress, the ER lipid composition is remodeled to involve an increased proportion of saturated phospholipids⁷⁶. Moreover, acyl chain saturation of membrane lipids is crucial for membrane fluidity, with unsaturated lipid acyl chains enabling better fluidity and saturated lipid acyl chains forming nonfluid and packed structures^77,78. Study has shown that increased membrane saturation/order leads to a loss of fluidity, which may affect the dynamic association and disassociation between organelles, further leading to an increase in MERCS⁴⁴. Therefore, we investigated the lipidome of the ER to assess the relationships among ER lipid composition, membrane fluidity, and ER–mitochondria contact, as well as the regulatory role of NM-NTUs in this process.

We identified 877 metabolites in the ER lipidome, with 392 significantly different metabolites between the IL-1β + NM-NTU + light group and the IL-1β + NM-NTU + dark group (Fig. 6a; Supplementary Fig. 13a). PCA revealed distinct clustering among samples within groups, with similar metabolite profiles in the control and IL-1β + NM-NTU + light groups, while both groups clustered separately from the IL-1β and IL-1β + NM-NTU + dark groups (Fig. 6b). Compared with the IL-1β + NM-NTU + dark group, the IL-1β + NM-NTU + light group significantly affected Ca²⁺-dependent events and lipid metabolism (Fig. 6c; Supplementary Fig. 13b). Kyoto Encyclopaedia of Genes and Genomes enrichment analysis revealed that NM-NTU + light treatment affected pathways related to the biosynthesis of unsaturated fatty acids (Supplementary Fig. 13c).

**Fig. 6: NM-NTUs remodel ER lipid composition.**

We subsequently analyzed the distribution and changes in ER lipid categories among the groups. Compared with those in the control group, the triglyceride (TG) levels were significantly reduced in the IL-1β and IL-1β + NM-NTU + dark groups, whereas they were restored in the IL-1β + NM-NTU + light treatment group (Fig. 6d). The levels of other major ER lipid species (e.g., phosphatidylcholine, phosphatidylethanolamines, phosphatidylinositol, and sphingomyelin) did not change significantly among the groups (Fig. 6e–h).

We further used thin-layer chromatography (TLC) to determine the TG content in the ER of each group. Compared with the IL-1β + NM-NTU + dark group, the IL-1β + NM-NTU + light group contained more TGs (Fig. 6i). To address whether ER–mitochondria contact was related to TG levels, we modulated TG levels in the ER by applying inhibitors of TG synthesis. The final and committed step of TG synthesis is catalyzed by two diacylglycerol acyltransferases, DGAT1 and DGAT2, which are ER-resident transmembrane enzymes⁷⁹. We treated NPCs with a DGAT1 inhibitor (A-922500) and a DGAT2 inhibitor (PF-06424439) or a combination of both. TLC analysis of ER lipids confirmed that inhibiting DGAT1 and DGAT2, either individually or in combination, reduced the TG levels in the ER (Fig. 6j). We then performed immunofluorescence experiments using calnexin and TOMM20 in cells treated with DGAT1 and DGAT2 inhibitors. Compared with the control group, the inhibition of TG synthesis significantly increased the number of MERCS (Fig. 6k, l). These results suggested a link between TG levels and the phenotype of MERCS. We further discuss the reasons for this link. Since inhibition of TG synthesis disrupts the channeling of fatty acids into neutral lipid storage⁸⁰ and disruption of lipid homeostasis elicits ER stress⁸¹, the formation of MERCS may increase as an adaptive response to the unfolded protein response^82,83. In general, our findings indicated that an altered ER lipidome (reduced TG levels) might contribute to increased ER–mitochondria contact.

We subsequently analyzed the acyl chain compositions of TG. Compared with those in the IL-1β + NM-NTU + dark group, the proportions of TG (54:3)[NL-18:1], TG (50:2)[NL-18:1], TG (54:2)[NL-18:0], TG (56:3)[NL-18:1], and TG (52:3)[NL-16:1] significantly increased in the IL-1β + NM-NTU + light group (Fig. 6m). The TG volcano map further confirmed these results (Fig. 6n). The proportion of unsaturated TG in the IL-1β + NM-NTU + light group was significantly greater than that in the IL-1β + NM-NTU + dark group (Fig. 6o). In summary, NM-NTU + light treatment increased the unsaturation levels of TGs. Given that lipid unsaturation significantly increased membrane fluidity⁷⁷ and affected the contact between intracellular organelles⁴⁴, we speculated that the NM-NTU reduced the formation of MERCS in degenerative cells by enhancing membrane fluidity. Next, we measured global membrane fluidity in NPCs using a membrane fluidity kit. Membrane fluidity was quantified by measuring the ratio of excimer to monomer fluorescence of lipophilic pyrene probes⁸⁴. With increasing membrane fluidity, the probe underwent excimer formation. We found that NPCs in the IL-1β + NM-NTU + light group exhibited greater membrane fluidity than did those in the IL-1β + NM-NTU + dark group (Fig. 6p). Overall, NM-NTUs with light illumination might affect ER–mitochondria contact by remodeling the lipid composition of the ER.

NM-NTUs ameliorate IVDD in rats

To further evaluate the in vivo efficacy of the NM-NTUs, we constructed a puncture-induced IVDD model in rats^85,86. Co7/8 intervertebral discs were selected for surgery (Fig. 7a). At 4 and 8 weeks post-surgery, micro-computed tomography (CT) radiographs showed that the intervertebral discs that underwent needle puncture without subsequent NM-NTU + light treatment had a low disc height index (DHI). The DHI of the NM-NTU + light treatment group was higher than that of the NM-NTU + dark group (Fig. 7b, c; Supplementary Fig. 14), indicating that the NM-NTU + light treatment maintained the DHI. DHI restoration is usually accompanied by increased disc water content. Magnetic resonance imaging (MRI) was used to assess disc water content in the different groups. The results showed that NM-NTU + light treatment significantly delayed progressive water loss (T2 signal intensity) in IVDD (Fig. 7d, e). We evaluated the histological changes in the intervertebral disc after surgery. The discs in the sham group showed intact and well-defined microstructures. Compared with that in the vehicle and NM-NTU + dark groups, NM-NTU + light treatment attenuated NP destruction at 4 and 8 weeks post-surgery (Fig. 7f, g). Alcian blue, safranin O-fast green, and Picrosirius red staining were used to further evaluate NP histological morphology (Fig. 7h–m). NP was replaced by fibrous tissues in the vehicle and NM-NTU + dark groups. In contrast, the NM-NTUs in the light group showed a normal NP appearance and a clear boundary between the NP and the annulus fibrosus. Fluorescence staining was performed to investigate the NM-NTU in vivo mechanism of action. Compared with the vehicle and NM-NTU + dark groups, the NM-NTU + light group showed higher Col II and aggrecan protein levels in the intervertebral discs (Fig. 8a, b). Similarly, ER stress-related proteins (AFT4 and CHOP) were detected. The results showed that NM-NTU + light treatment reduced AFT4 and CHOP levels compared with those in the vehicle and NM-NTU + dark groups (Fig. 8a, b). We subsequently conducted immunofluorescence costaining of calnexin and TOMM20 in rat IVD tissue sections. The results indicated that compared with the sham group, MERCS increased in the puncture + vehicle and puncture + vehicle + NM-NTU + dark group. Moreover, MERCS significantly decreased in the NM-NTU + light treatment group compared with the NM-NTU + dark treatment group (Fig. 8c, d), which revealed that the abnormal contact between the ER and mitochondria in rat IVDD was attenuated. We further extracted cells from the NP tissue and performed TEM to further evaluate the MERCS in the tissue samples. For each field of view, we systematically quantified the ER tubules that were closely apposed to the mitochondria, the number of mitochondria, and the average length of the ER tubules. Interestingly, the NM-NTU + light group showed a significant reduction of ER closely apposed to mitochondria compared with the puncture + vehicle and puncture + NM-NTU + dark group (Fig. 8e, f). ER length and mitochondrial counts were calculated for each field, and the results revealed no significant difference in the number of mitochondria or the average length of ER tubules among the four groups (Fig. 8g, h). These results provided partial evidence that the in vivo therapeutic effects of NM-NTUs were functionally linked to their ability to restore the organelle interaction network. In addition, histological staining of multiple organs indicated NM-NTU biosafety in vivo (Supplementary Fig. 15). Taken together, these results reveal that NM-NTUs have therapeutic effects in rat IVDD models.

**Fig. 8: Effects of NM-NTUs on protein levels and MERCS in vivo.**

Remote wireless controllable device for deep illumination

Although NM-NTU has shown therapeutic potential in rats, its application to the intervertebral discs of larger animals may be limited by poor light penetration. Therefore, developing a safe and implantable light device for NTU treatment is crucial. The development of wireless power technology facilitates light delivery to deep tissues through wireless devices⁸⁷. First, we assembled the wp-LED by connecting the LED parallel to the inductor and capacitor (Fig. 9a; Supplementary Fig. 16). Various color lighting was achieved using wireless charging coils (Fig. 9b). To enable remote control of the wp-LED, we connected the wireless coil to a smart socket, which served as the receiver. Hence, we controlled the receiver using a mobile phone (via a wireless network) and remotely controlled the wp-LED switch (Supplementary Movie 1). In addition, the lighting duration was precisely controlled using a mobile phone (Supplementary Movie 2). As wp-LEDs need to be implanted in the body, an insulating, biocompatible, and nondegradable transparent packaging material is required. Polyurethane (PU) materials have been used for the insulation packaging of cardiac implantable electronic devices because of their good biocompatibility and non-degradability⁸⁸. Therefore, PU was used to package the wp-LED (PU-wp-LED), minimizing the risk of inflammation and device rejection. The PU-wp-LED was immersed in a phosphate buffer solution to confirm whether it could be insulated in a wet environment. The results showed the effectiveness of PU encapsulation, as the wp-LED remained functional and emitted light (Fig. 9c). Subsequently, we assessed the wp-LED output efficiency. In the horizontal plane, the closer to the center of the coil, the lower the brightness of the wp-LED, according to the principle of electromagnetic induction. In the vertical plane, the wp-LED light intensity decreased with increasing distance from the coil center (Fig. 9d). The PU and PU-wp-LED leachates also exhibited no evident cytotoxicity (Fig. 9e). We subcutaneously implanted the PU-wp-LEDs to test their feasibility in animals. Upon activating wireless power via the mobile phone, the device effectively emitted red light from the rabbit subcutaneous tissue (Fig. 9f; Supplementary Movie 3). Similarly, the results of rabbit body temperature and blood tests showed that the PU-wp-LED exhibited good biosafety (the early postoperative temperature increase may have been caused by surgery) (Fig. 9g; Supplementary Table 1). Furthermore, we simulated deep intervertebral disc illumination by surgically implanting a PU-wp-LED ventral to the disc and injecting NM-NTUs into the disc using X-ray fluoroscopy (Fig. 9h, i). The CT and X-ray results confirmed fixation of the PU-wp-LED ventral to the intervertebral disc and the feasibility of intradiscal NM-NTU injection (Fig. 9i, j). Moreover, we simulated the use of PU-wp-LED on the human body, in which the device was fixed to the human forearm. When the mobile power supply was turned on, the red light was effectively emitted, confirming the potential application value of this device (Supplementary Fig. 17). Overall, we established a therapeutic approach for treating deep degenerative tissues using wireless-powered light combined with NTUs.

PU-wp-LED combined with NM-NTUs alleviate IVDD in rabbits

To determine whether the light from the PU-wp-LED had sufficient intensity to activate NM-NTUs, we first determined that NM-NTUs had significant therapeutic effects in cell experiments with light intensities above 10.3 μmol photons m⁻² s⁻¹ (Supplementary Fig. 18a). This light intensity was defined as the light threshold for the activation of NM-NTUs. Furthermore, in the in vivo rabbit photoirradiation experiment, we measured the distance from the back skin of the intervertebral disc to the anterior longitudinal ligament by MRI, which was 4.45 ± 0.36 cm (Supplementary Fig. 18b). We then estimated that the PU-wp-LED output power in the rabbit body (intervertebral disc area) within the coil was approximately 60–70 μmol photons m^-2 s^-1. Considering the approximately 60% red light transmission efficiency through the annulus fibrosus (Supplementary Fig. 18c), we estimated that the light intensity reaching the nucleus pulposus was above the activation threshold of NM-NTUs. We further verified the therapeutic effect of PU-wp-LED combined with NM-NTUs in the rabbit IVDD model (Fig. 10a). At 4 and 8 weeks post-surgery, MRI results showed the signal intensity of discs in the NM-NTU + PU-wp-LED + light group reduced less compared to the dark group (Fig. 10b, c). Then we obtained fresh disc tissues and observed their appearance. The discs in the vehicle group and NM-NTU + PU-wp-LED + dark group both showed severe degeneration, while the NM-NTU + PU-wp-LED + light group showed mild changes compared with sham group (Fig. 10d). Moreover, compared to the dark group, the NM-NTU + PU-wp-LED + light group attenuated NP degeneration in histological analysis (Fig. 10e–h). We also detect the protein levels of Col II and aggrecan in different groups. The NM-NTU + PU-wp-LED + light treatment elevated both Col II and aggrecan levels compared to the dark group (Fig. 10i). We also conducted experiments in the rabbit IVDD model, comparing PU-wp-LED and external light (Supplementary Fig. 19a). Compared with the NM-NTU + external light group, MRI results revealed that discs treated with the NM-NTU + PU-wp-LED + light maintained significantly greater signal intensity, indicating better water content and reduced disc dehydration (Supplementary Fig. 19b, c). Histological evaluation via H&E staining further supported these findings. In the NM-NTU + PU-wp-LED + light group, the overall disc morphology appeared well preserved compared with that in the NM-NTU + external light group (Supplementary Fig. 19d). Histological scoring based on an established grading system confirmed a significant improvement in the NM-NTU + PU-wp-LED + light group compared with the NM-NTU + external light group (Supplementary Fig. 19e). Overall, compared with direct external light, the PU-wp-LED system achieved favorable therapeutic outcomes in the rabbit IVDD model. The reason for the external light system failure might be its inability to sufficiently penetrate dorsal anatomical structures and reach deep tissues. Taken together, the remote controllable wireless irradiation device combined with NM-NTU effectively alleviated deep disc degeneration in rabbits.

In summary, organelle interaction networks can become dysfunctional under pathological conditions, such as degeneration. Implanting plant-derived independent photosynthetic systems into mammalian cells can effectively restore homeostasis of the interaction network of animal organelles, potentially owing to cellular energy supply by NTUs. Furthermore, NTUs can regulate intracellular Ca²⁺ homeostasis and membrane fluidity by increasing ER lipid unsaturation and reducing MERCS formation and mitochondrial dysfunction. By combining implantable LEDs with wireless charging technology, we expanded photosynthesis therapy, enabling smarter and more effective treatments for deeper tissues. This treatment model has broad application prospects for various diseases involving imbalanced organelle interaction networks.

Methods

Cell culture

Rat and human NPCs were obtained from NP tissues by collagenase digestion for 4 h⁸⁹. Rat AFCs were obtained from annulus fibrosus tissues by collagenase and hyaluronidase digestion for 1.5 h⁹⁰. Rat EPCs were obtained from endplate cartilage tissues by collagenase digestion for 6 h⁹¹. Rat NP tissues, annulus fibrosus tissues and endplate cartilage tissues were macroscopically isolated from 4-week-old male Sprague–Dawley rats. Rat NPCs, AFCs, and EPCs were seeded with media changed every 72 h, passaged when the confluence reached 80%, and utilized before passage 2. Human NP tissues were obtained from human participants undergoing spinal surgery for vertebral fractures or IVDD. Cultures of primary human NPCs were maintained under standard conditions (5% CO₂ and 37 °C), and the medium was changed every 72 h. The human NPCs were passaged on the sixth day. To evaluate the impact of in vitro culture on the phenotypic changes in degenerated human NPCs, cell samples were collected every other day from day 1 to day 9 of culture. These samples were subsequently used for gene and protein analysis of NPC phenotypic markers. The use of patient samples was approved by the Ethics Committee of Sir Run Run Shaw Hospital. Informed consent was obtained from all the patients involved in the study. The cells were seeded in a humidified incubator and cultured in Dulbecco’s modified Eagle medium (DMEM), supplemented with 10% fetal bovine serum (Gibco). For plasmid transfection, we used IH-NPCs (iCell Bioscience, Inc., iCell-0028a), which were cultured in specialized culture medium (iCell Bioscience, Inc.).

NTU and NM-NTU preparation

Chloroplasts were isolated from fresh spinach leaves using differential centrifugation and Percoll density gradient centrifugation, and then suspended in a hypotonic lysis buffer to obtain thylakoids^19,92. The thylakoids were extruded sequentially through polycarbonate membranes with pore sizes of 800-, 400-, 200-, and 100-nm to obtain NTUs. The NPC membranes were isolated from disrupted NPCs by differential centrifugation¹⁹ and suspended in water at a concentration of 2 mg mL⁻¹. Subsequently, the NPC membrane solution was mixed with an equal volume of the NTUs and allowed to stand for 30 min. Next, the mixed solution was filtered through 1000-, 400-, and 200-nm polycarbonate membranes using an Avanti mini extruder. NM-NTUs were obtained through centrifugation at 10,000 × g for 5 min and were resuspended for further use. The chlorophyll content was determined by spectrophotometric analysis⁹³. To evaluate the encapsulation efficiency of the NTUs by NM, we used nanoFCM. NTUs were stained with StarRed and detected in the PC5 channel. The NMs were labeled with a FITC-conjugated marker, allowing their specific detection in the FITC channel. Different NTU and NM protein ratios (ranging from 1:10 to 1:200) were tested. Events that were positive for both FITC and PC5 (FITC⁺/PC5⁺) were classified as NMs successfully loaded with NTUs, whereas events that were positive for FITC but negative for PC5 (FITC⁺/PC5^-) were categorized as empty NMs. Events that were positive for PC5 but negative for FITC (FITC^-/PC5⁺) were categorized as free NTUs. NTU and NM-NTU morphologies were observed using cryo-TEM (200 kV, FEI Tecnai G2 F20). The size and surface zeta potential of the nanoparticles were determined via dynamic light scattering using a Malvern Zetasizer Nano ZS49. SDS-PAGE and Coomassie Blue (Beyotime) staining were used to detect proteins in the thylakoids, NTUs, NPC membranes, and NM-NTUs.

Antibodies

The following antibodies were used for western blotting: CHOP (Proteintech, 15204-1-AP, 1:1,500); IRE1 (Proteintech, 27528-1-AP, 1:1,500); ATF6 (Proteintech, 66563-1-Ig, 1:10,000); ATF4 (Proteintech, 60035-1-Ig, 1:2000)；β-actin (Proteintech, 20536-1-AP, 1:5000).

For immunofluorescence analysis, the following antibodies were used: KRT19 (Proteintech, 10712-1-AP, 1:200), CD24 (Proteintech, 10600-1-AP, 1:200), CHOP (Proteintech, 15204-1-AP, 1:100); ATF4 (Proteintech, 60035-1-Ig, 1:200); Col II (Proteintech, 28459-1-AP, 1:800); aggrecan (Proteintech, 13880-1-AP, 1:200); MMP13 (Proteintech, 18165-1-AP, 1:200); ADAMTS-5 (Affinity Biosciences, DF13268, 1:200); TOMM20 (Abcam, ab56783, 1:200); Calnexin (Proteintech, 10427-2-AP, 1:200).

Cell viability assay

Human NPC viability was evaluated using two independent assays: the MTT assay (Beyotime) and the live/dead staining assay (Beyotime). An MTT assay was performed to measure cellular metabolic activity. Briefly, yellow, water-soluble MTT tetrazolium salt was added to the cell culture. The absorbance at 570 nm was measured spectrophotometrically. A live/dead staining assay was used to simultaneously distinguish viable cells from nonviable cells on the basis of membrane integrity. Cells were stained with a combination of calcein AM and propidium iodide. Images were acquired using a laser confocal microscope (LCM, Nikon), and the percentages of live and dead cells were quantified using Fiji/ImageJ software. Rat NPCs were incubated with NTU or NM-NTUs (0.5 μg mL⁻¹ for chlorophyll) for 24 h. A CCK-8 assay was performed to determine cell viability (Dojindo).

Cellular uptake of NM-NTUs

We evaluated the cellular uptake of the NM-NTUs by rat AFCs, EPCs and NPCs. Briefly, cells were incubated with a medium containing NM-NTUs (DiI labeled NTUs, 0.5 μg mL⁻¹ for chlorophyll) for 6 h, followed by washing, fixation with 4% formalin, and DAPI staining. The cells were imaged and analyzed using an LCM (Nikon). NPCs were cultured, labeled with DiI, and treated with a medium containing NM-NTUs (DiO labeled NM, 0.5 μg mL⁻¹ for chlorophyll) for 1 h to detect the membrane fusion ability of NM-NTUs. Samples were fixed with 4% formalin, stained with DAPI, and photographed using an LCM (Nikon).

Intracellular trafficking of NTUs in NPCs

NTUs or NM-NTUs (DiI labeled NTUs, 0.5 μg mL⁻¹ for chlorophyll) were co-cultured with rat NPCs for 6 h, and lysosomes were labeled with lysoTracker Green (Beyotime) to investigate the co-localization of NTUs and lysosomes in cells. Subsequently, cells were fixed with 4% formalin and stained with DAPI; images were captured using an LCM (Nikon). Next, Mander’s overlap coefficients of lysosomes and NTUs were analyzed using Fiji/ImageJ (JACoP plugin). Colocalization studies were performed to determine the intracellular distribution of internalized NM-NTUs. After rat NPCs were incubated with NM-NTUs^DiI (NTUs labeled with DiI) for 6 h, the samples were stained for specific cellular structures, including the endoplasmic reticulum (ER-Tracker Green, Beyotime), mitochondria (Mito-Tracker Green, Beyotime), Golgi apparatus (Golgi-Tracker Green, Beyotime), actin filaments (Actin-Tracker Green, Beyotime), and microtubules (Tubulin-Tracker Green, Beyotime). Confocal microscopy images were acquired, and colocalization was quantitatively analyzed using Mander’s overlap coefficients. The Mander’s coefficients (M1) quantify the fraction of the signal intensity of NTUs that colocalizes with lysosomes relative to the total signal intensity of NTUs.

To visualize MERCS, IH-NPCs were transfected with SPLICS Mt-ER Short P2A (Addgene plasmid # 164108) encoding the mitochondrial outer membrane-targeted OMM-GFP1-10 and the endoplasmic reticulum-targeted short protein GFPβ11 (1 μg plasmid of a 35 mm glass bottom dish) using Lipofectamine 3000, strictly following the manufacturer’s protocol. At 36–48 h post-transfection, the cells were treated with DiI-labeled NM-NTUs for 6 h. After treatment, the cells were imaged using confocal microscopy. The reconstituted GFP signal, which indicates close-range (~8–10 nm) MERCS, and the DiI signal (NTUs) were captured. Colocalization between NTUs and MERCS puncta was quantitatively analyzed using the JACoP plugin in Fiji/ImageJ software, with the results expressed as the Mander’s coefficient (M1) representing the fraction of the NTU signal overlapping with MERCS.

Effects of NM-NTUs on cell metabolism

Rat NPCs were treated with IL-1β (20 ng mL⁻¹) for 24 h and incubated with NM-NTUs (0.5 μg mL⁻¹ for chlorophyll) for 6 h, with or without red light irradiation (from light-emitting diodes, peaking at 630 nm at an intensity of 80 µmol photons m⁻² s⁻¹, 30 min). The NPCs were incubated with the corresponding antibodies for immunofluorescence staining. Images were captured using an LCM (Nikon, Tokyo, Japan). Images were analyzed and fluorescence intensity was measured using the Fiji/ImageJ software. The intracellular ATP concentration was analyzed using an ATP assay kit (Beyotime). To evaluate the ATP production efficiency of the NM-NTU at different concentrations, we measured ATP levels under NM-NTU concentrations ranging from 0.125 to 2 μg mL⁻¹ for chlorophyll. Similarly, ATP production under varying light conditions (10.3–108.1 µmol photons m⁻² s⁻¹) was also assessed. To specifically quantify ATP production from NM-NTUs, we inhibited mitochondrial oxidative phosphorylation and glycolysis using rotenone (500 nM) and 2-DG (10 mM). After the NM-NTUs were subjected to light treatment for 30 min, ATP levels were measured using an ATP assay kit. For intracellular NADPH quantification, cells from each group were lysed and analyzed using an NADPH assay kit (Beyotime) according to the manufacturer’s instructions.

Assessment of endogenous mitochondrial functional recovery

IH-NPCs were transfected with pCMV-Mito-AT1.03 (Beyotime, D2606). Cells were seeded in 6-well plates, and each well was transfected with 1 µg of plasmid using Lipofectamine 3000. At 24–36 h post-transfection, the cells were treated with IL-1β for an additional 24 h. Mitochondrial ATP levels were subsequently quantified by flow cytometry on the basis of the ratio of YFP to CFP fluorescence following NM-NTUs + light treatment.

Quantitative polymerase chain reaction (qPCR)

Total RNA was extracted from the lysed NPCs using the SteadyPure RNA Extraction Kit (Accurate Biology). RNA was reverse-transcribed into cDNA using Evo M-MLV RT Master Mix (Accurate Biology), and qPCR was performed using the Hieff UNICON qPCR SYBR Green Master Mix on a QuantStudio^TM 6 Flex Real-Time PCR System (Thermo Fisher Scientific). Target gene sequences are listed in Supplementary Table 2.

Determination of cytoplasmic and mitochondrial Ca²⁺ concentration

NPCs were loaded with 2 μM Rhod-2/AM (50 min; Yeasen). The cells were then washed and reloaded with 5 μM Fluo-4/AM (30 min; Beyotime)⁹⁴. Images were recorded using an LCM (Nikon). Image analysis and fluorescence intensity measurements were performed using the Fiji/ImageJ software.

Detection of ER Ca²⁺ concentration

To investigate the contribution of photosynthetic ATP to ER calcium storage, IH-NPCs were transfected with pCMV G-CEPIA1er (Addgene# 58215), which encodes an ER calcium indicator⁶⁵. Photosynthetic ATP production in the NM-NTU was inhibited by treatment with 10 μM DCMU. The experimental design included eight treatment groups analyzed at 0, 15, and 30-min time points: control, IL-1β, IL-1β + NM-NTU + dark, and IL-1β + NM-NTU + light, with each condition further divided into subgroups either treated with or without DCMU. Cells were imaged using an LCM (Nikon), and quantitative analysis of fluorescence intensity was performed using Fiji/ImageJ software to assess [Ca²⁺]_ER.

mPTP evaluation

The mPTP of the NPCs was evaluated based on the calcein release assay⁹⁵ using an mPTP assay kit (Beyotime) and photographed using an LCM (Nikon).

MERCS observation

MERCS structures of the NPCs were observed using TEM (H-9500; Hitachi). MERCS in TEM images were manually analyzed in Fiji/ImageJ. Briefly, the image scale was first set with the Set Scale tool, and the freehand line tool was subsequently used to measure the length of the ER or membrane or the perimeter of the mitochondria. The contact length was defined as the ER membrane length (within 0–100 nm of the mitochondria) divided by the mitochondrial perimeter. Mitochondria and ER architecture in the TEM images were reconstructed graphically using Adobe Illustrator software. Similarly, the MERCS structure of the NPCs was observed by immunostaining the ER and mitochondria with calnexin and TOMM20. To quantify MERCS, we performed colocalization analysis using Fiji/ImageJ plugins (Coloc2 and JACoP) following established methods⁹⁶. The Mander’s coefficients (M1) quantify the percentage of the signal intensity of mitochondria that colocalizes with the ER relative to the total signal intensity of the mitochondria.

Membrane fluidity detection

The global membrane fluidity of the NPCs was evaluated using a membrane fluidity kit (Abcam), which utilizes fluorescent lipophilic pyrene probes that are incorporated into the cell membrane. The monomeric pyrene probe undergoes excimer formation, shifting its emission from Em 400 nm to Em 470 nm (excitation at 360 nm). Membrane fluidity was quantified by determining the ratio of excimer (Em 470 nm) to monomer (Em 400 nm) fluorescence using a fluorescence microplate reader (BioTek).

Intracellular ROS and mitochondrial membrane potential detection

A ROS assay kit (Beyotime) was used to detect intracellular ROS. The DCFH-DA (Beyotime) fluorescence emission was acquired by flow cytometry or LCM. Furthermore, a JC-1 fluorescent probe (Solarbio) was used to detect mitochondrial membrane potential in NPCs. Image analysis and fluorescence intensity measurement were performed using the Fiji/ImageJ software.

Western blot analysis

NPCs were lysed using RIPA buffer (FDbio). SDS-PAGE gels (FDbio) were used to separate the proteins. Proteins were transferred onto polyvinylidene difluoride membranes (FDbio) after electrophoresis. After blocking with skim milk (FDbio) and incubation with the primary antibody, bands were developed using Horseradish peroxidase-labeled secondary antibodies (Beyotime). Following chemiluminescence detection, bands were visualized using the ChemiDoc Touch imaging system (Bio-Rad).

Transcriptomic and lipidomic study

Human NP tissues and rat NPCs were used for transcriptomic analysis. Briefly, total RNA was extracted using TRIzol reagent, following the manufacturer’s instructions. Sequencing was performed using an Illumina NovaSeq 6000 following the manufacturer’s protocol recommended by the supplier. Genes with a P value < 0.05 and an absolute log₂(fold changes) ≥ 1 were considered differentially expressed. GSEA was performed using GSEA software (https://www.broadinstitute.org/gsea/). Volcano maps and other visual analyses were performed using the OmicStudio tools (https://www.omicstudio.cn/tool).

For the lipidomic experiment, the ER from rat NPCs was isolated using an ER isolation kit (Sigma), and sample pretreatment and analysis were conducted. Data analysis and acquisition were primarily performed using a Vanquish Flex UHPLC system (Thermo Fisher Scientific) and a TSQ Altis Plus mass spectrometer (Thermo Fisher Scientific). Quantification of lipid species was determined by comparison to the relevant internal standard using Skyline 22.2.0527. All detected lipids were annotated and classified based on the lipid databases (https://www.lipidmaps.org/; http://lipidontology.com/). Metabolites with P < 0.05 and fold change > 10% were considered significant.

TLC analysis

Thin-layer chromatography (TLC) was used for TG quantification⁹⁷. Specifically, the total lipids in the ER extracts were obtained by suspending the ER in 1 mL of PBS supplemented with 0.1 mM phenylmethylsulphonyl fluoride (PMSF) and 2 mL of chloroform/methanol (2:1 vol/vol) supplemented with 0.01% butylated hydroxytoluene. After centrifugation at 1500 × g for 5 min, the organic phases were collected and dried with nitrogen and resuspended in chloroform. TLC was performed by spotting the total lipid extract dissolved in chloroform onto 10 cm EMD TLC Silica Gel plates (Labshark) and developing them with hexane/diethyl ether/acetic acid (80:20:2, vol/vol). Lipids were visualized with iodine vapor prior to imaging.

PU-wp-LED assembly

The 220 pF multi-layer ceramic capacitor (221K, 0603) and surface-mount-device (SMD) LED (0805) were soldered on a 2.2 MH SMD power inductor (222, CD43) in parallel. Subsequently, wp-LED was encapsulated in PU and wrapped with poly-wires. The PU-wp-LEDs were detected immediately after assembly using a wireless charging coil (XINKETAI Electronic Co., Ltd.). The light intensity of the device was measured using a spectral illuminance meter (Linshang LS330). A laser power meter (Zhongxi Equipment) was used to measure the efficiency of red light penetrating the annulus fibrosus (3 mm thickness) from the sheep.

Biosafety of PU-wp-LED

To determine the biosafety of PU or PU-wp-LED in vitro, we soaked PU or PU-wp-LED in DMEM for 48 h to obtain the leachate. Subsequently, the NPCs were cultured with the leachate for 24 h. CCK-8 assay (Dojindo) was used to detect cell viability.

Animal models

The animal use protocol was reviewed and approved by the Institutional Animal Care and Use Committee of the Zhejiang Center of Laboratory Animals. Male Sprague–Dawley rats (12 weeks old) were used for in vivo experiments. The IVDD rat model was established using the needle puncture method⁹⁸. First, rats were anesthetized with pentobarbital sodium. Subsequently, the IVDD models were established using a 26 G needle puncture of the coccyx spaces Co7–8. The needle reached the NP center and was removed after 30 s. Rats were randomly divided into four groups, namely sham surgery (sham), puncture and 5 µL vehicle (Puncture + Vehicle) injection per week, puncture + 5 µL NM-NTUs (20 μg mL⁻¹ for chlorophyll) injection per week without red light irradiation (Puncture + NM-NTUs + dark), and puncture + 5 µL NM-NTU injection (20 μg mL⁻¹ for chlorophyll) per week with red light irradiation for 30 min daily (Puncture + NM-NTUs + light). At week 4 and 8, the rats were euthanized for further evaluation.

Male New Zealand rabbits (6–7 months old) were used for in vivo experiments. The rabbits were anesthetized with pentobarbital sodium and placed in a prone position on a platform. The PU-wp-LEDs were implanted subcutaneously to assess their safety and lighting performance. Briefly, a small incision was made in the dorsal skin of the rabbit, and the subcutaneous tissue was separated using a blunt probe; subsequently, the PU-wp-LED was inserted subcutaneously, and the wound was sutured (PU-wp-LED group). The illumination capability of the PU-wp-LED in vivo was tested immediately after surgery. The sham group underwent a surgical procedure similar to that of the implantation group; however, it did not involve PU-wp-LED implantation (sham group). The body temperature of the rabbits was measured daily for 1 week after implantation, and routine blood tests were performed 2 weeks after implantation. For the intervertebral disc illumination simulation, rabbit spines were exposed using a lateral retroperitoneal approach with aseptic techniques. Next, PU-wp-LED was implanted anterior to the L4/5 disc, fixed to the soft tissue structure with a silk thread, and the incision was closed. The intervertebral disc was punctured using a 20 G needle under X-ray guidance to simulate deep NM-NTU injection.

For rabbit IVDD treatment, the rabbits were anesthetized and their spines were exposed. L4/5 IVD were punctured with an 18 G needle at a depth of 5 mm, then PU-wp-LED was implanted anterior to the L4/5 disc and fixed to the soft tissue structure with silk thread, and the incision was closed. The following intra-disc injection was via X-ray fluoroscopy guide. Rabbits were randomly divided into 4 groups: sham surgery (sham), puncture and 10 µL vehicle (Puncture + Vehicle) injection per week, puncture + 10 µL NM-NTUs (20 μg mL⁻¹ for chlorophyll) injection per week with PU-wp-LED turn off (Puncture + PU-wp-LED + NM-NTUs + dark), and puncture + 10 µL NM-NTU injection (20 μg mL⁻¹ for chlorophyll) per week with PU-wp-LED turn on for 30 min daily (Puncture + PU-wp-LED + NM-NTUs + light). At week 4 and 8, the rabbits were euthanized for further evaluation. And PU-wp-LED were removed before imaging evaluation. To compare the effects of implantable versus nonimplantable optical stimulation, the rabbits were randomly assigned to one of four groups: sham surgery (sham), puncture and 10 µL vehicle (Puncture + Vehicle) injection per week, puncture + 10 µL NM-NTUs (20 μg mL⁻¹ for chlorophyll) injection per week with external red light illumination for 30 min daily (Puncture + NM-NTUs + External illumination), and puncture + 10 μL NM-NTU injection (20 μg mL⁻¹ for chlorophyll) per week with PU-wp-LED turn on for 30 min daily (Puncture + NM-NTUs + PU-wp-LED illumination). The PU-wp-LED implants were removed prior to imaging evaluation. All the animals were euthanized at week 8 for subsequent analysis.

In vivo radiological assessment

Magnetic resonance imaging was performed on vertebral samples 4 and 8 weeks postoperatively using a GE Sigma CV/I (1.5 T). The rat caudal vertebral samples were scanned using micro-CT (Skyscan1275). We defined the regions of interest to cover the upper and lower normal intervertebral spaces. The calculation method for DHI is shown in Supplementary Fig. 14. DHI was calculated using the following equation:

$${{{\rm{DHI}}}}=2\times ({{{\rm{A}}}}+{{{\rm{B}}}}+{{{\rm{C}}}})/({{{\rm{D}}}}+{{{\rm{E}}}}+{{{\rm{F}}}}+{{{\rm{G}}}}+{{{\rm{H}}}}+{{{\rm{I}}}})$$

(1)

where A, B, and C represent the disc height between adjacent vertebrae; and D, E, F, G, H, and I represent the length of the vertebral bone immediately rostral and caudal to the intervertebral disc. Changes in the DHI were expressed as a percentage (%DHI = measured DHI/Sham group DHI × 100). Fiji/ImageJ software was used to measure the signal intensity on the T2-weighted central images. A CT scan was used to confirm the fixation of the PU-wp-LED anterior to the rabbit intervertebral disc.

Histological analysis

Disc samples were fixed, decalcified, and embedded to obtain the sections. The sections were stained with hematoxylin and eosin (H&E), Alcian blue, Safranin O-fast green, or Picrosirius red. The disc samples were scored according to the table shown in Supplementary Table 3. Subsequently, the sections were incubated with the corresponding antibodies for immunofluorescence staining. Rat major organs were collected at 8 weeks for systemic toxicity assessment (through H&E staining).

Statistics and reproducibility

Data were presented as mean ± standard deviations. The statistical analysis methods were described in the figure legends. Unpaired two-tailed Student’s or Welch’s t test (two independent groups) and one-way analysis of variance with Tukey’s post hoc test (three or more groups) were used. Ordinal grading system data were analyzed using the Kruskal–Wallis and post-hoc Dunn’s tests. Statistical significance was set at P < 0.05. Statistical analyses were performed using Excel and GraphPad Prism software version 10.0. Particle size analysis, zeta potential, SDS-PAGE, cryo-TEM, Western blot, and in vitro fluorescence imaging experiments were independently repeated three times. H&E staining of major organs in rats were replicated in two independent experiments.

Reporting summary

Further information on research design is available in the Nature Portfolio Reporting Summary linked to this article.

Data availability

Transcriptomics data are available at NCBI BioProject under accession PRJNA1405942 and PRJNA1405940. The lipidomics data generated in this study have been deposited in the MetaboLights database with dataset identifier MTBLS13730. Source data are provided with this paper.

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Acknowledgements

This work was supported by grants from the National Nature Science Fund of China Grant (grant numbers 82372454 and T2422022 to P.C., 82322043 to X.L., and 82330077 to S.F.), Scientific Research Innovation Capability Support Project for Young Faculty (ZYGXQNJSKYCXNLZCXM-H19 to X.L.), the New Cornerstone Science Foundation through the XPLORER PRIZE (to X.L.), Natural Science Foundation of Zhejiang Province (grant numbers LR24H060001 to P.C. and TGY24H060025 to C.X.), China National Postdoctoral Program for Innovative Talents under (grant number BX20240327 to C.G.), Ningbo Major S&T Project (grant number 2024Z211 to P.C.) and the Zhejiang Provincial Program for the cultivation of high-level innovative health talents (to P.C.).

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These authors contributed equally: Chen Xia, Zhanqiu Dai, Yongcheng Wang.

Authors and Affiliations

Department of Orthopaedic Surgery, Sir Run Run Shaw Hospital, Zhejiang University School of Medicine, Hangzhou, China
Chen Xia, Zhanqiu Dai, Yongcheng Wang, Jiachen Yu, Zijie Wang, Zhenxiang Huang, Kaifeng Pan, Ke Yang, Yiyu Chen, Peiyu Zhong, Chenhui Gu, Shunwu Fan, Xianfeng Lin & Pengfei Chen
Center for Plastic and Reconstructive Surgery, Department of Orthopedics, Zhejiang Provincial People’s Hospital, Affiliated People’s Hospital Hangzhou Medical College, Hangzhou, China
Chen Xia
Key Laboratory of Mechanism Research and Precision Repair of Orthopaedic Trauma and Aging Diseases of Zhejiang Province, Hangzhou, China
Chen Xia, Zhanqiu Dai, Yongcheng Wang, Jiachen Yu, Zijie Wang, Zhenxiang Huang, Kaifeng Pan, Ke Yang, Yiyu Chen, Peiyu Zhong, Chenhui Gu, Shunwu Fan, Xianfeng Lin & Pengfei Chen
Department of Orthopaedics, The Second Affiliated Hospital and Yuying Children’s Hospital of Wenzhou Medical University, Wenzhou, China
Zhanqiu Dai
Cixi Biomedical Research Institute, Wenzhou Medical University, Ningbo, China
Xianfeng Lin & Pengfei Chen

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Chen Xia
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Conceptualization: P.C., X.L., S.F., C.X., and C.G. Data curation: C.X., Z.D., and Y.W. Formal analysis: C.X., Z.D., Y.W., J.Y., and Z.W. Funding acquisition: P.C., X.L., S.F., C.X., and C.G. Investigation: C.X., Z.D., Y.W., J.Y., Z.W., Z.H., K.P., K.Y., Y.C., and P.Z. Methodology: P.C., C.X., Z.D., and Y.W. Project administration: P.C. Supervision: P.C., X.L., S.F., and C.G. Validation: P.C. C.X., Z.D., and Y.W. Visualization: C.X. and Z.D. Writing original draft: C.X. and Z.D. Writing, review and editing: P.C., X.L., S.F., and C.G. All authors have read and approved the article.

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Correspondence to Chenhui Gu, Shunwu Fan, Xianfeng Lin or Pengfei Chen.

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Nature Communications thanks Maria Ankarcrona, who co-reviewed with Ming Ho Choi; Cao Yang and the other, anonymous, reviewer(s) for their contribution to the peer review of this work. A peer review file is available.

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Xia, C., Dai, Z., Wang, Y. et al. Natural photosynthetic system for restoring homeostasis of animal organelle interaction network. Nat Commun 17, 3087 (2026). https://doi.org/10.1038/s41467-026-69825-y

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Received: 19 April 2025
Accepted: 10 February 2026
Published: 31 March 2026
Version of record: 31 March 2026
DOI: https://doi.org/10.1038/s41467-026-69825-y