COG5 deficiency disrupts cellular copper homeostasis and underlies the impaired mitochondrial OXPHOS function
Yuwei Zhou, Keyi Li, Ruowei Zhu, Xue Ma, Xinfei Ye, Mengqing Mao, Ding Li, Xiaofei Zeng, Zhehui Chen, Jing Wu, Liqin Jin, Xiaohua Tang, Yanling Yang, Jianxin Lyu, Xiaoting Lou, Carmen Priolo, Carmen Priolo, Carmen Priolo, Carmen Priolo
TL;DR
COG5 deficiency causes copper overload and mitochondrial dysfunction, leading to energy production defects and a new form of Leigh syndrome.
Contribution
Identifies COG5's role in copper homeostasis and mitochondrial function, revealing a novel pathogenic mechanism and potential therapeutic target.
Findings
COG5 deficiency disrupts ATP7A stability, leading to cellular copper overload.
Excess copper impairs mitochondrial iron-sulfur clusters and complex I assembly.
Copper chelation rescues mitochondrial dysfunction in COG5-deficient models.
Abstract
COG5, a subunit of the conserved oligomeric Golgi (COG) complex, plays a critical role in retrograde trafficking within the Golgi apparatus. Dysfunction of COG5 is associated with various human disorders, yet the underlying pathogenic mechanisms remain poorly understood. To investigate the mechanisms, we conducted proteomic analyses using COG5-deficient and rescue cell models, which revealed a potential link between COG5 dysfunction and mitochondrial oxidative phosphorylation (OXPHOS) deficiency. Using COG5-deficient cell models and patient-derived cells harboring COG5 variants, we biochemically validated the involvement of COG5 in mitochondrial OXPHOS, particularly in the regulation of complex I content. These models also exhibited elevated cellular copper levels. Notably, the significant reduction in OXPHOS complexes could be rescued by either restoring COG5 expression or…
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Fig 6- —http://dx.doi.org/10.13039/501100001809National Natural Science Foundation of China
- —http://dx.doi.org/10.13039/501100001809National Natural Science Foundation of China
- —http://dx.doi.org/10.13039/501100004731Natural Science Foundation of Zhejiang Province
- —http://dx.doi.org/10.13039/501100004731Natural Science Foundation of Zhejiang Province
- —Zhejiang Provincial Medical and Health Science and Technology Plan Project
- —Key Discipline of Zhejiang Province in Medical Technology (First Class, Category A), Wenzhou Medical University
- —Key Discipline of Zhejiang Province in Public Health and Preventive Medicine (First Class, Category A), Hangzhou Medical College.
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TopicsMitochondrial Function and Pathology · Alzheimer's disease research and treatments · Cellular transport and secretion
Introduction
The conserved oligomeric Golgi (COG) complex is a critical regulator of vesicular trafficking within the Golgi apparatus, ensuring the proper sorting and transport of proteins and lipids necessary for maintaining Golgi structure and function [1]. COG complex consists of eight subunit proteins divided into two subassemblies: COG1, COG2, COG3, and COG4 in lobe A and COG5, COG6, COG7, and COG8 in lobe B [2–7], which are highly conserved evolutionarily and present in all eukaryotes [4,8,9]. Among its subunits, COG5 has emerged as a key player in mediating retrograde trafficking, a process essential for Golgi homeostasis [5,10]. Despite its fundamental role in intracellular trafficking [11,12], the molecular mechanisms underlying COG5 dysfunction and its contribution to human diseases remain poorly understood. To date, only 19 cases of COG5 deficiency have been reported, all of which are associated with a spectrum of multisystemic disorders, including developmental delay, hypotonia, ataxia, epilepsy, leukodystrophy, and ichthyosis [13–20]. However, the precise pathogenic pathways linking COG5 deficiency to cellular dysfunction and the resulting disease phenotypes remain largely unexplored. This knowledge gap underscores the urgent need for further research to elucidate the molecular and cellular consequences of COG5 impairment.
Copper, an essential metal for all known eukaryotes, plays a vital role in numerous cellular processes, including mitochondrial function and iron-sulfur cluster (ISC) biogenesis. Intriguingly, recent studies have revealed that COG5 interacts with ATP7A [21,22], a copper-transporting ATPase [23], suggesting a potential role for COG5 in copper homeostasis. Disruption of copper homeostasis has been shown to impair mitochondrial function [24–26], raising the possibility that mitochondrial oxidative phosphorylation (OXPHOS) deficiency may contribute to the pathogenesis of COG5-related diseases. However, the mechanistic link between COG5 deficiency, copper dysregulation, and mitochondrial dysfunction remains unclear. A comprehensive investigation into the molecular pathways underlying COG5-related diseases is therefore needed.
In this study, we aimed to address this critical gap by investigating whether COG5 deficiency-induced copper dysregulation disrupts ISC stabilization, thereby contributing to OXPHOS dysfunction. Using COG5-related cell models and patient-derived cells carrying biallelic COG5 variants, we sought to elucidate the molecular mechanisms connecting COG5 dysfunction to mitochondrial impairment and disease pathogenesis. Our findings provide new insights into the role of COG5 in cellular homeostasis and offer a potential framework for understanding the broader implications of COG5 deficiency in human diseases. A graphical abstract summarizing the key findings of this study is provided as Fig 1.
COG5 deficiency disrupts copper homeostasis, impairing mitochondrial OXPHOS functions.Loss of COG5 disrupts cellular copper homeostasis, leading to a deficit in the stabilization of iron-sulfur clusters (ISCs). The impairment in ISC biogenesis directly compromises the function of mitochondrial OXPHOS complexes, ultimately resulting in mitochondrial dysfunction and the manifestation of disease phenotypes. Created in BioRender. Yuwei, Z. (2025) https://BioRender.com/w0nfm7v and Yuwei, Z. (2025) https://BioRender.com/t4sc9bl.
Results
Proteomics analysis implicates mitochondrial dysfunction, oxidative stress, and energy metabolism as potential mechanisms underlying COG5 deficiency.
To determine the potential mechanisms underlying the COG5 deficiency, proteomics analysis was performed based on cell models generated in HEK293T cells: Control, COG5-knockout (KOs) and rescue model cells (KO + COG5), and the modulation of COG5 protein levels were confirmed by western blot (Fig 2A and S1A Fig). The schematic diagram of the proteomics analysis process is shown in Fig 2B and S1 Data. Principal component analysis (PCA) separated the proteomic profiles of the Control, KO1, and KO1 + COG5 groups, indicating significant differences in their biological characteristics (Fig 2C). The ranking of proteins based on relative abundance in KO1 compared to the Control revealed differential expression levels of COG-related proteins, with the eight subunits of the COG complex containing proteins noted in Fig 2D. A cartoon diagram of the COG complex, where the relative expression of single subunits was indicated by color intensity, demonstrated that depleting COG5 reduced the abundance of all lobe-B subunits, particularly COG7, consistent with previous research [9] (Fig 2E). The chord diagram summarized the differential expression between KO1 and the Control, besides the N-glycan biosynthesis pathways which are already known [27–30], surprisingly oxidative phosphorylation was most highlighted (Fig 2F). Collectively, these results suggest that COG5 plays a significant role in mitochondrial function and cellular metabolism which needs further verification.
COG5 knockout disrupts COG complex integrity and mitochondrial protein expression.A. Western blot analysis of the steady-state levels of COG5 in HEK293T cells featuring KO1, rescue cells (KO1 + COG5), and corresponding control cells (Control). Control cells were obtained by transfecting empty control vectors. β-Actin served as an internal control. B. Schematic illustrating the proteomic analysis workflow in HEK293T cells featuring KO1, rescue cells (KO1 + COG5), and corresponding control cells (Control). Control cells were generated by transfecting empty control vectors. Details of the experimental groupings are provided. Created in BioRender. Yuwei, Z. (2025) https://BioRender.com/na67l1k. C. Principal component analysis (PCA) of global proteomes in HEK293T-based cell models with KO1, rescue cells (KO1 + COG5), and matched control cells (Control). Each point represents an independent biological replicate. D. Overlay of the differentially expressed proteins in KO1 compared with Control, with COG complex subunits (COG1-COG8) were explicitly highlighted. E. Schematic representation of the conserved oligomeric Golgi (COG) complex architecture. Subunits are depicted as circles, with the color intensity corresponding to their relative protein abundance in KO1 compared to Control (Left), and in rescue (KO1 + COG5) compared KO1 (Right). Lobe A (COG1-4) and lobe B (COG5-8) are annotated. F. This chord diagram showing the results of proteomic analysis in KO1 cells compared with Control. Differentially expressed proteins are clustered into five distinct biological pathways. Proteins localized to the mitochondrial oxidative phosphorylation complexes are specifically highlighted in purple.
COG5 restoration rescue impaired OXPHOS function in COG5 KO cells
To further investigate the relationship between COG5 and mitochondrial OXPHOS function, we isolated and quantified respiratory chain complexes using blue-native PAGE. We found that complex I (CI) and complex III_2_ (CIII_2_) in COG5 KO cells were significantly reduced than paired control cells. The re-expression of COG5 restored the contents of CI and CIII_2_ further verified the correlation of COG5 deficiency and OXPHOS complex deficiency (Fig 3A-3B and S1B-C Fig). Furthermore, COG5-deficient cells exhibited defects in the activities of multiple mitochondrial respiratory enzymes, particularly CI and CIII₂, which aligns with the blue-native PAGE results (Fig 3C and S1D Fig). Schematic representation of mitochondrial complexes subunits along its structure modules first described (Fig 3D), and the heatmap further summarizes the changes in protein abundance within the mitochondrial complexes (Fig 3E). Finally, the COG5 KO cells displayed reduced cellular ATP levels and elevated mitochondrial superoxide production compared to control cells and were restored by re-expression of COG5 (Fig 3F-3G and S1E-S1F Fig). To assess the ultrastructural consequences of COG5 deficiency, we performed transmission electron microscopy (TEM). This analysis revealed severe abnormalities in both the Golgi apparatus and mitochondria in KO cells compared to Control. The Golgi stacks appeared fragmented and dilated, with numerous swollen vacuoles replacing the characteristic compact cisternal organization. Concurrently, mitochondria were markedly swollen and displayed disorganized and fragmented cristae. These morphological alterations provide clear evidence that COG5 loss disrupts the structural integrity of key cellular organelles (S1G Fig). Collectively, we consider that COG5 can affect OXPHOS, especially the content of CI.
*Proteomic analysis indicates that COG5 deficiency results in impaired oxidative phosphorylation.A-B. Analysis of steady-state levels of OXPHOS complexes in HEK293T cells with KO1, KO1 + COG5, and paired control cells (A), quantified as shown in (B). 2% Triton X-100 solubilized cells were separated and visualized by 3.5%-16% blue-native PAGE. The OXPHOS complexes were detected by immunoblotting with antibodies against proteins as indicated (CI, Grim19; CII, SDHA; CIII2, UQCRC2, CIV, MTCO1; CV, ATP5A), with TOM40 serving as the internal loading control. C. Enzymatic activities of mitochondrial electron transport chain (ETC) complexes (CI-CIV) in KO1, KO1 + COG5, and control cells. D. Schematic representation of mitochondrial OXPHOS complexes and their subunit composition, highlighting proteins identified in proteomic analysis. E. Heatmap showing expression changes of mitochondrial OXPHOS complex in KO1 compared to Control based on proteomic analysis. F. Whole-cellular ATP production in HEK293T cells with KO1, KO1 + COG5, and their paired control cells was normalized to protein concentration. G. Detection of mitochondrial superoxide production in HEK293T cells with KO1, KO1 + COG5, and their paired control cells. Quantitative data from three independent experiments were analyzed and presented as the means ± SEM. ns, Statistical significance is denoted as ns, no significance, *p < 0.05, **p < 0.01, ***p < 0.001, ***p < 0.0001.
COG5 deficiency induced intracellular copper overload and impaired respiratory chain complexes levels
Given the essential role of copper in mitochondrial function and the established association between COG5 and ATP7A [22,31], we postulated that COG5 regulates OXPHOS through modulation of cellular copper homeostasis. This proposed mechanism is illustrated in the schematic diagram depicting COG5-mediated mitochondrial dysfunction (Fig 4A). We first asked whether COG5 interacts with ATP7A to facilitate its stability. Co-immunoprecipitation assays confirmed a physical interaction between COG5 and ATP7A (Fig 4B). Consequently, the loss of COG5 led to a significant reduction in ATP7A protein levels reduced by approximately 50%, as validated by both Western blot (Fig 4C-4D) and proteomic analysis (Fig 4E) and, without affecting its mRNA expression (Fig 4F), suggesting a post-translational mechanism. To directly test if COG5 regulates the turnover of ATP7A protein, we performed a cycloheximide (CHX) chase assay. The results demonstrated that ATP7A was degraded significantly faster in KO cells compared to Control (Fig 4G), confirming that COG5 binding facilitates ATP7A protein stability. We next hypothesized that the collapse of ATP7A protein levels impairs copper export, leading to intracellular accumulation. Indeed, ELISA analysis revealed a significant increase in cellular copper content in KO cells, of which were rescued by COG5 re-expression (Fig 4H).
*Redundancy of intracellular copper reduces the expression level of OXPHOS complexes.A. Schematic model of ATP7A-mediated copper transport under physiological conditions and under copper overload caused by COG5 deficiency. Under normal conditions, ATP7A traffics between the trans-Golgi network (TGN) and plasma membrane to maintain copper homeostasis. Disruption of COG5 may impair ATP7A localization and function, leading to copper retention and mitochondrial toxicity. Created in BioRender. Yuwei, Z. (2025) https://BioRender.com/na67l1k. B. HEK293T cells were co-transfected with a plasmid encoding COG5-Flag. Whole-cell lysates were immunoprecipitated using antibodies against ATP7A and COG5, or a control mouse IgG. The immunoprecipitates along with the input lysates (representing 5% of total lysate) were analyzed by Western blotting with the indicated antibodies. C-D. Western blot analysis of ATP7A protein expression in Control, KOs and rescue model cells (C), quantified as shown in (D). Whole-cell lysates were probed with antibodies against ATP7A and β-Actin. β-Actin served as an internal control. E. Schematic representation of the COG complex and ATP7A. Circle color intensity reflects relative protein abundance of COG subunits (COG1–COG8) and ATP7A in KO1 compared to Control (Left), and in rescue (KO1 + COG5) compared to KO1 (Right), based on proteomic analysis. F. Quantitative RT-PCR (qRT-PCR) analysis of ATP7A mRNA levels in Control, KOs and rescue model cells (KO + COG5). mRNA expression was normalized to β-Actin and is presented as fold change relative to the control group. G. ATP7A protein stability levels at indicated time points (0, 3, 6, 9 h) in Control and KO1 after 100 μM cycloheximide (CHX) were analyzed by Western Blot. β-Actin served as an internal control. H. Intracellular copper content was determined by ELISA in Control, KOs and rescue model cells (KO + COG5) based on HEK293T cells. Copper content in cells was normalized to total protein concentration. I-J. Control cells were treated with 1 μM CuCl₂ and 0.17 μM disulfiram in the culture medium for 12 hours. Triton X-100-solubilized lysates were separated on a 3.5%–16% gradient gel with the indicated antibodies. Quantification of the assembled OXPHOS complexes as shown in (J). TOM40 was used as a loading control. K-L. Blue-native PAGE (3.5%-16%) analysis of OXPHOS complexes using triton-solubilized cell lysate with the indicated antibodies in KO1 and Control cells treated with 20 μM tetrathiomolybdate (TTM) solubilized in culture medium for 4 hours and were quantified as in (L). TOM40 was used as an internal loading control. Quantitative data from three independent experiments were analyzed and presented as the means ± SEM. ns, Statistical significance is denoted as ns, no significance, *p < 0.05, **p < 0.01, ***p < 0.001, ***p < 0.0001.
We subsequently investigated whether copper overload could affect the expression of OXPHOS complexes. As reported, previous studies mainly focused on the effect of copper-underloading on OXPHOS, for instance, mitochondrial complex IV is a crucial copper-dependent enzyme with two copper-binding sites in its catalytic core, the reduced level of copper level caused a deficiency of mitochondrial function and respiratory-chain complex biosynthesis, especially CIV [32–34]. Although the effects of copper deficiency on OXPHOS have been extensively studied [32], the impact of the expression of holistic OXPHOS complexes affected by overload copper is rarely still unknown. To address this, we employed an established cellular paradigm of copper overload, treating HEK293T cells with 1 μM CuCl₂ in combination with the copper ionophore disulfiram (0.17 μM). This copper concentration regimen, adapted from the work of Tsvetkov et al. [35], is a well-validated model for inducing intracellular copper accumulation to pathologically relevant levels. As demonstrated in Fig 4I-4J, exposure to elevated copper resulted in a significant reduction in the levels of both CI and CIII₂ compared to untreated controls, confirming that copper availability is a critical regulator of mitochondrial OXPHOS complex expression.
To determine whether modulation of copper transport could rescue the OXPHOS defects, we exogenously expressed ATP7A in KO1 cells. The overexpression levels of ATP7A was confirmed by both qRT–PCR and Western blotting analyses (S2A-S2C Fig). Notably, ATP7A expression alleviated the intracellular copper accumulation observed in KO1 cells (S2D Fig). Consistent with improved copper homeostasis, the abundance of OXPHOS complexes was partially restored following ATP7A overexpression (S2E Fig). These results suggest that impaired copper export contributes to mitochondrial dysfunction in COG5-deficient cells and that restoring ATP7A expression mitigate these defects. These findings demonstrate that COG5 deficiency destabilizes ATP7A leading to intracellular copper overload.
We next proposed that the reduced expression of CI and CIII₂ in COG5 KO cells results from copper overload. To validate this hypothesis, we employed tetrathiomolybdate (TTM), a specific copper chelator, to modulate intracellular copper levels.
To determine an appropriate working concentration of TTM while minimizing cytotoxicity, Control and KO1 cells were exposed to increasing TTM concentrations across multiple time points. Cell number analysis indicated that lower concentrations (≤2 μM) exerted minimal effects on cell viability, whereas higher concentrations or prolonged exposure led to a progressive decline in cell numbers in both groups, indicating dose- and time-dependent cytotoxicity (S3A-S3B Fig). These data defined an initial tolerability range for subsequent functional studies. To further evaluate whether copper chelation could rescue mitochondrial defects, a short-term dose-response analysis was performed focusing on OXPHOS complex abundance. This analysis demonstrated a concentration-dependent restoration of OXPHOS complexes, with maximal recovery observed following short-term exposure to 20 μM TTM for 4 h (S3C Fig), suggesting that transient higher-dose treatment can achieve functional rescue despite partial cytotoxic effects observed during longer exposures. Meanwhile, we also conducted the treatment with a conservative low concentration. Blue native PAGE analysis showed that treatment with 2 μM TTM for 12 h partially restored OXPHOS complex abundance in KO1 cells (S3D Fig). Based on the balance between rescue efficacy and acceptable short-term toxicity, 20 μM TTM for 4 h was therefore selected as the optimal condition for subsequent experiments. TTM treatment resulted in significant restoration of CI and CIII₂ expression in COG5 KO cells. As anticipated, complex IV expression was markedly reduced in both control and COG5 KO cells following TTM treatment [36], while CII and CV levels remained unchanged (Fig 4K-4L and S4A-S4B Fig). These results provide compelling evidence that COG5 deficiency-induced copper overload specifically impairs the expression of key OXPHOS complexes, particularly CI and CIII₂. Collectively, these results suggest that superfluous copper caused by COG5-deficient reduces the expression level of OXPHOS complexes.
COG5 deficiency impairs CI through disrupting ISC activity
Our previous observations revealed a substantial reduction in CI levels in COG5-deficient cells. Given the displacement of native iron-sulfur clusters (ISCs) centers by excess Cu [37] and the crucial role of ISCs in CI biogenesis [38,39]. We investigate whether COG5 deficiency impacts ISC activity. Firstly, we performed a comprehensive re-analysis of ISC-related proteins from proteomic data, followed by heatmap analysis categorizing proteins based on their localization and functional roles, including OXPHOS subunits, ISC assembly factors, and cytosolic iron-sulfur cluster assembly (CIA) components. The results demonstrated a significant reduction in ISC-containing OXPHOS subunits, particularly those associated with CI (Fig 5A). Furthermore, separation of mitochondrial respiratory super-complexes by Blue-native PAGE revealed more maturely assembled mitochondrial super-complexes exhibit assembly defects, as evidenced by the accumulation of lower molecular weight sub-assemblies (S4C Fig).
*Loss of COG5 causes an abnormal iron-sulfur cluster.A. Heatmap illustrating the expression of proteins based on the classification of iron-sulfur proteins based on proteomics. Proteins are grouped by molecular function, including Fe-S assembly factors (ISC and CIA machinery) and Fe-S containing subunits of OXPHOS complexes. B-C. Western blot (WB) analysis of the steady-state levels of ACO-2 in COG5 KO cells (KO1, KO2) and paired control cells. The expression levels were quantified as in (C). D-E. In gel enzymatic activities assay of ACO-2 in mitochondrial extracts from KO1, KO2 and paired control cells. The activity levels were quantified as in (E) expressed as relative values of Control activity. F-G. In gel enzymatic activities of ACO-2 in mitochondrial extracts from KO1, KO1 + OE, and Control cells. The activity levels were quantified as in (G). H-I. Measurement of cellular iron-sulfur cluster content using a fluorescent biosensor in KO1, KO1 + COG5, and control cells. The contents were quantified as in (I). J. Proposed model illustrating how COG5 deficiency leads to copper accumulation, which disrupts Fe-S cluster biogenesis or stability, resulting in defective assembly and function of Complex I—an Fe-S cluster-rich complex I. Created in BioRender. Yuwei, Z. (2025) https://BioRender.com/xm0hlaq. Quantitative data from three independent experiments were analyzed and presented as the means ± SEM. ns, Statistical significance is denoted as ns, no significance, *p < 0.05, **p < 0.01, ***p < 0.001, ***p < 0.0001.
We next examined aconitase (ACO-2), a mitochondrial ISC-containing enzyme. While mitochondrial ACO-2 protein levels remained unchanged in COG5 KO cells compared to controls (Fig 5B-5C). However, the results showed that COG5 KO cells showed decrease in mitochondrial ACO-2 in-gel activity compared with control cells (Fig 5D-5E). Recover with exogenous COG5, the activity of ISC was restored in recovery cell models (Fig 5F-5G). At the same time, we also detected the content of ISC in whole-cell to rule out the effect of content production on activity. The results showed that COG5 KO cells showed no significant decrease in mitochondrial ACO-2 content (Fig 5H-5I). Collectively, mitochondrial ISC functional activity is lost in COG5 KO cells, and we speculate that this is one of the mechanisms responsible for the defective CI assembly (Fig 5J).
Clinical presentations and pathogenicity evaluation of COG5 variants
As previously described, genetic-based cell models have demonstrated that COG5 deficiency leads to copper overload and impaired ISC activity, resulting in mitochondrial OXPHOS dysfunction. We hypothesized that COG5 deficiency might exist in mitochondrial disease patients. Here, we searched the in-house database of clinically diagnosed mitochondrial disease patients without known disease-causing genes. We identified a patient carrying biallelic COG5 variants who clinically presented with mitochondrial disease for further analysis.
First, we reviewed the medical records as follows. The proband (II-1), a 12-year-old boy, was born at term as the first child of non-consanguineous Chinese parents following an uncomplicated pregnancy and spontaneous vaginal delivery. At 2 years and 10 months, he was referred to our pediatric clinic due to sudden mental-motor regression, manifested by an inability to walk, difficulty standing, tremors and weakness in the upper limbs, hypertonia in the lower limbs, speech impairment, limited expressiveness, excessive fatigue, poor digestion, occasional coughing, and irritability. Ophthalmic examination revealed bilateral hyperopia and concomitant left eye strabismus. His anthropometric measurements include a BMI of 17.54 kg/m^2^ (the range of +1SD to +2SD), height of 95.5 cm, weight of 16 kg, and head circumference of 50.2 cm (the range of median to +1SD). At 3 years and 7 months, his anthropometric measurements include a BMI of 18.14 kg/m^2^ (the range of +1SD to +2SD), height of 105 cm, weight of 20 kg, cranial perimeter of 49.5 cm (the range of -1SD to median). At 7 years and 11 months, his anthropometric measurements include a BMI of 20.83 kg/m^2^ (the range of +2SD to +3SD), height of 120 cm, weight of 30 kg, cranial perimeter: 52.2 cm.
Laboratory examination revealed an elevated blood lactate level, with 3.49 mM (reference range: 0.50-2.20 mM), without abnormalities in blood amino acids, acylcarnitine, urine organic acids tests, and lysosomal storage diseases enzymatic analysis. Brain magnetic resonance imaging (MRI) showed asymmetric abnormal signals in the bilateral basal ganglia, brainstem, and cerebellar hemispheres. These were mainly manifested as symmetrical small patches with slightly low signal on T1-weighted imaging (T1WI), slightly high signal on T2-weighted imaging (T2WI), and T2-fluid-attenuated inversion recovery (FLAIR) in both basal ganglia regions and cerebellar hemispheres (S5A-S5D Fig). A summary of the clinical features and detailed clinical case histories are provided in S5E Fig and Table 1.
Table 1: Genetic and clinical features of COG5 patients.
The whole exome sequencing (WES) and mitochondrial DNA sequence was performed with the patient blood sample, according to the criteria mentioned before [40], two biallelic variants in COG5 (NM_006348.5) c.1290C > A (NP_006339.4: p. Phe430Leu) and c.1826T > C (NP_006339.4: p. Ile609Thr) were identified and then verified by sanger sequence (Fig 6A). As shown in S6A Fig, the allele frequency of c.1290C > A and c.1826T > C in gnomAD_exomes was extremely low (0.0003015, 0.0003893), in gnomAD_genomes was extremely low (0.0000691, 0.000779), in 1000 genomes and ExAc was also low (0.00260, 0.001353 and 0.0032, 0.001776). The pathogenicity of c.1290C > A and c.1826T > C was predicted by silico analysis (SIFT, PROVEAN, and MutationTaster), resulting that c.1290C > A might be damaging, neutral, and disease-causing and c.1826T > C might be damaging, deleterious, and disease-causing, respectively. Taken together, c.1290C > A and c.1826T > C might disease-causing variants. Both p.F430 and p.I609 were highly conserved among species suggesting they are important for the function of COG5 protein (S6B Fig). Both variants were predicted to be deleterious by multiple in silico tools and are located in evolutionarily conserved regions. Notably, these variants were present in population databases at low allele frequencies, and homozygous individuals were reported, which initially challenged their potential pathogenicity. To functionally assess their impact, we established patient-derived lymphoblastoid cell lines and site-directed mutagenesis cell models harboring each variant (Mut 1: COG5 ^c.1290C>A^, Mut 2: COG5 ^c.1826T>C^). Western blot analysis revealed a significant reduction in COG5 protein expression in both patient-derived and site-directed mutagenesis cell models (Fig 6B–6E). In summary, despite the population frequency data, our functional assays provide compelling evidence for a deleterious effect of these variants, supporting their potential contribution to the patient’s phenotype.
*Analysis of COG5 amount and mitochondrial OXPHOS functions in patient-derived immortalized lymphocytes.A. The biallelic COG5 variants in the patient (c.1290C > A and c.1826T > C) were identified by the Sanger sequencing. B-C. Western blot (WB) analysis of the steady-state levels of COG5 in immortalized lymphocytes from the patient (P) and paired healthy controls (C1, C2), and the abundances of blots were quantified as in (C). β-Actin was used as an internal loading control. D-E. Western blot (WB) analysis of the steady-state levels of COG5 in mut cell models (mut 1: c.1290C > A and mut 2: c.1826T > C) and paired controls were quantified as in (E). β-Actin was used as an internal loading control. F-G. Blue-native PAGE (3.5%-16%) and immunoblotting analysis of the OXPHOS complexes in patient-derived immortalized lymphocytes (P) and paired healthy controls (C1, C2) and the abundances of blots were quantified as in (G). Antibodies against CI (Grim19), CII (SDHA), CIII2, (UQCRC2), CIV (MTCO1), and CV (ATP5A) were used to detect OXPHOS complexes. TOM40 was used as an internal loading control. H. Cellular ATP production in immortalized lymphocytes from the patient (P) and healthy controls (C1, C2) was normalized to cell numbers. I. Production of mitochondrial superoxide in immortalized lymphocytes from the patient (P) and healthy controls (C1, C2). J-K. Western blot analysis of ATP7A protein expression in in immortalized lymphocytes from the patient (P) and healthy controls (C1, C2) (J), quantified as shown in (K). Whole-cell lysates were probed with antibodies against ATP7A and β-Actin. β-Actin served as an internal control. L. Intracellular copper content was determined by ELISA in immortalized lymphocytes from the patient (P) and healthy controls (C1, C2). Copper content in cells was normalized to total protein concentration. M-N. Blue-native PAGE (3.5%-16%) and immunoblotting analysis of the OXPHOS complexes in patient-derived immortalized lymphocytes (P) and paired healthy controls (C1, C2) and were quantified as in (N). Antibodies against CI (Grim19), CII (SDHA), CIII2, (UQCRC2), CIV (MTCO1), and CV (ATP5A) were used to detect OXPHOS complexes. TOM40 was used as an internal loading control. -: do not any treatment; + : module cells were treated with 20 μM tetrathiomolybdate (TTM) solubilized in culture medium for 4 hours. Quantitative data from three independent experiments were analyzed and presented as the means ± SEM. ns, Statistical significance is denoted as ns, no significance, *p < 0.05, **p < 0.01, ***p < 0.001, ***p < 0.0001.
Patient-derived lymphocytes with a decreased COG5 level and a deficit in mitochondrial OXPHOS function regulated by copper content
To verify the pathogenic role of COG5 deficiency in mitochondrial OXPHOS dysfunction, we performed further biochemical analyses using patient-derived lymphocytes. Blue-native PAGE analysis revealed a significant reduction in CI expression of COG5 deficit patient cells compared to controls (Fig 6F-6G). Consistently, a significant reduction in whole-cell ATP content was observed in patient cells (Fig 6H) accompanied by elevated mitochondrial superoxide levels (Fig 6I), indicating compromised mitochondrial bioenergetics and increased oxidative stress.
As mentioned earlier, copper homeostasis is closely linked to mitochondrial function in COG5 KO based on HEK293T cells, we further examined the copper transporter ATP7A patient lymphocyte. Western blot analysis showed decreased ATP7A protein expression in patient lymphocytes relative to healthy controls (Fig 6J–6K). Consistent with this observation, intracellular copper content was significantly increased in patient cells (Fig 6L), suggesting disrupted cellular copper handling in COG5-deficient lymphocytes. Similarly, we treated patient-derived lymphocytes with 20 μM TTM. Remarkably, TTM treatment restored CI expression to near-normal levels (Fig 6M-6N), supporting our hypothesis of copper-mediated OXPHOS dysfunction.
These findings collectively demonstrate that COG5 deficiency in patient-derived lymphocytes is associated with reduced OXPHOS capacity and mitochondrial dysfunction, which are mechanistically linked to copper dysregulation. The restoration of CI expression following copper chelation suggests a potential therapeutic strategy for mitigating mitochondrial dysfunction in COG5-deficient conditions.
Summary of previously reported patients carrying the COG5 variants
In addition, dysfunctional COG complex leads to defective cellular glycosylation modifications, which can clinically lead to CDG. CDG resulting from COG complex defects could be referred to as COG-CDG. The effect of COG complex defects on glycosylation was first described in 1986 [5], and the first case of CDG due to defects in COG7 was reported in 2004 [41]. At present, 19 patients carrying with variants in COG5 have been reported from 12 reported studies. We summarized the reported variants in COG5 (S7 Fig) and clinical manifestations of the previously reported patients (Table 1 and S2 Data).
Discussion
In eukaryotes, the transport of proteins and lipids among intracellular compartments is mediated by vesicular and tubular carriers under the direction of an elaborate protein machinery. MTCs are thought to mediate the initial attachment (or tethering) between a trafficking vesicle and its target membrane through a constellation of interactions; of which, COG5, one subunit of COG complex, is an MTC that is essential for vesicle transport within the Golgi apparatus and from endosomal compartments to the Golgi [27–30]. COG5 was recently identified as an interactome gene for ATP7A implicated in neurodegeneration and neurodevelopmental disorders [21]. Faundez [22] demonstrated that COG complex activity is required for mitochondrial respiration. Given the established role in glycosylation within the COG complex, we hypothesized that COG5, and by extension the COG complex, might regulate mitochondrial function through mechanisms beyond its established role in glycosylation.. In this study, we demonstrate that COG5 plays an essential role in preserving CI integrity and mitochondrial function by regulating cellular copper homeostasis, as evidenced in both cellular models and patient-derived cells.
To investigate the potential role of COG5 in mitochondrial OXPHOS regulation, we built upon previous reports of an interaction between COG5 and the major copper transporter ATP7A [22]. We also identified that (1) a physical interaction between COG5 and ATP7A confirmed by co-immunoprecipitation (Fig 4B); (2) unaltered ATP7A mRNA levels (Fig 4F); and (3) markedly decreased ATP7A protein stability upon COG5 loss (Fig 4C-4E). Based on these findings and the established role of the COG complex in Golgi retrograde trafficking, we propose that COG5 is essential for the proper localization and stability of ATP7A. In the absence of COG5, ATP7A recycling may be disrupted, leading to its mis-localization and subsequent degradation via cellular quality control pathways. We further hypothesized that COG5 may function as a regulator of copper homeostasis, with downstream implications for mitochondrial respiration. ATP7A plays a central role in cellular copper homeostasis, yet the consequences of its dysfunction are highly context-dependent. At the organismal level, ATP7A deficiency classically leads to systemic copper deficiency, as exemplified by Menkes disease, where impaired export of dietary copper from intestinal epithelial cells into the circulation results in reduced copper delivery to peripheral tissues. In contrast, at the cellular level ATP7A primarily mediates copper efflux and supplies copper to the secretory pathway via the trans-Golgi network; therefore, its loss can lead to intracellular copper accumulation rather than depletion. Indeed, copper retention in the cytosol, nucleus, and mitochondria has been reported in ATP7A-deficient cells [42], suggesting that disrupted copper export rather than impaired uptake predominates at the cell scale. Collectively, these observations highlight that the impact of ATP7A dysfunction on copper balance depends on biological scale, cell type, and membrane trafficking context, and should therefore be interpreted cautiously when linking ATP7A alterations to cellular copper status. Our results demonstrate that COG5 deficiency leads to significant intracellular copper accumulation, indicative of impaired copper distribution or export. Importantly, this defect was functionally rescued upon exogenous COG5 expression, as reflected by normalized copper levels and partial recovery of OXPHOS capacity. To further substantiate the role of COG5 in copper homeostasis, we extended our functional validation to AML-12 cells—a hepatocyte line recognized for its well-defined copper metabolic pathways. Consistent with our observations in HEK293T cells, COG5 knockdown (S8A Fig) in AML-12 cells resulted in a reduction in ATP7A protein levels (S8B-S8D Fig). This replication of the phenotype in a physiologically relevant cell model strengthens the conclusion that COG5-dependent regulation of ATP7A stability is not cell-type restricted but rather represents a general mechanistic principle. Importantly, this finding reinforces the central thesis of our study that COG5 acts as a critical regulator of cellular copper homeostasis.
Furthermore, extending our investigation beyond COG5, we found that knockdown of other COG complex subunits also led to reductions in the levels of CI and CIII_2_ (S9A-S9G Fig). This observation suggests that the regulation of mitochondrial OXPHOS integrity is not a function unique to COG5 but may be a common consequence of disrupting the COG complex. It reinforces the model that a functionally intact COG complex is essential for maintaining cellular copper homeostasis and, by extension, mitochondrial function. Clinically, an analysis of 39 reported COG-CDG cases (S3 Data) revealed frequent manifestations overlapping with MD spectra, such as developmental delay, deafness, epilepsy, and hepatic involvement. Nevertheless, detailed biochemical assessments of mitochondrial function—such as OXPHOS complex activities or systematic lactate/pyruvate analyses—are seldom available in clinical reports. The current reported COG-CDG patients’ clinical data are insufficient to definitively establish mitochondrial dysfunction as a consistent feature in these individuals.
Having established the link between COG5 loss and copper overload, we next sought to determine how this metabolic imbalance impairs mitochondrial function. Because mitochondria are the primary site of ISC biogenesis—a process essential for CI assembly—we hypothesized that elevated copper levels could interfere with mitochondrial ISC activity. Consistent with this possibility, ACO-2 protein levels remained unchanged in COG5-KO cells, whereas its enzymatic activity was reduced, suggesting impaired ISC functionality. These findings indicate that copper-mediated ISC disruption may contribute to the OXPHOS defects observed in COG5-deficient cells. The preservation of ACO-2 protein abundance alongside reduced activity further implies that copper excess preferentially affects ISC-dependent enzymatic function rather than causing generalized protein degradation. In addition, defects in the assembly of mature mitochondrial super-complexes were observed in COG5-deficient cells, as reflected by the accumulation of lower molecular weight sub-assemblies. (S4C Fig).
Our comparative analysis revealed distinct OXPHOS complex deficiency profiles between HEK293T cells and patient-derived lymphocytes (Figs 3A and 6F). Based on these observations, we hypothesized that differential dependence on COG5-mediated glycosylation might underlie these phenotypic variations. We treated both cell types with N-glycosylation inhibitors and observed cell line-specific patterns of complex deficiencies (S10A-S10B Fig). These findings suggest that: (1) The impact of COG5 on OXPHOS complexes is mediated, at least in part, through its role in protein glycosylation; (2) Different cell types possess varying degrees of reliance on COG5-dependent glycosylation for proper respiratory chain assembly.
The relationship between impaired ISC function, elevated mitoSOX, and OXPHOS deficiency in COG-deficient cells is likely multifactorial. Although ISC proteins are highly sensitive to oxidative damage and increased ROS could, in principle, contribute to reduced ISC enzyme activity, our data argue against ROS being the primary driver of the mitochondrial OXPHOS defects observed here. Pharmacological attenuation of ROS using N-acetylcysteine effectively lowered cellular oxidative stress but did not substantially restore OXPHOS complex abundance (S11A-S11B Fig). These findings suggest that mitochondrial dysfunction in COG-deficient cells is not simply a consequence of oxidative stress but is more directly linked to disrupted copper homeostasis, with elevated ROS likely representing a downstream consequence of respiratory chain imbalance rather than its initiating cause.
Diagnosis of rare diseases, especially multi-system involved including a pronounced neurologic damage, has always been a challenge in clinical. The development and application of NGS have provided considerable advancement in determining the molecular pathogenesis of heterogeneous diseases such as MD. The panel-based NGS strategy is a quick pathway to obtain a genetic diagnosis, but the effectiveness of this approach depends on the completeness of the correlation between clinical phenotype and genotype to ensure that the actual pathogenic gene is included in the panel. Therefore, expanding the pathogenic gene spectrum and clinical phenotype of diseases is essential [43–45]. In this study, based on the clinical cohort of MD, we filtered the WES sequencing data to discover novel pathogenic genes of MD and further explore the pathogenic mechanism [46]. Through comprehensive whole-exome sequencing analysis of our MD cohort, we identified a diagnostically challenging case exhibiting classical MD features without molecular confirmation. Systematic re-analysis of sequencing data revealed biallelic pathogenic variants in COG5, previously unreported in MD, establishing a novel genotype-phenotype correlation. The two COG5 variants (c.1290C > A [p. Phe430Leu] and c.1826T > C [p. Ile609Thr]) identified in the presented patient present a challenging scenario for clinical interpretation. The interpretation of the two identified COG5 variants (c.1290C > A [p. Phe430Leu] and c.1826T > C [p. Ile609Thr]) is complicated by conflicting evidence. Although their allele frequencies are low, the presence of homozygous individuals in population databases constitutes strong evidence for benign impact (BS2 according to ACMG/AMP guidelines). This stands in direct conflict with our functional studies, which provide strong evidence for pathogenicity (PS3) by demonstrating a clear deleterious effect on COG5 protein stability and cellular morphology. Specifically, patient-derived cells and isogenic models harboring these variants exhibited significantly reduced COG5 protein levels. Furthermore, ultrastructural analysis revealed Golgi fragmentation and abnormal mitochondrial cristae, supporting a loss of COG5 function. According to ClinGen recommendations, well-validated functional data can provide strong evidence for pathogenicity (PS3). An additional consideration arising from the genetic analysis of our patient is the possibility of intragenic (or allelic) complementation. While each variant has been reported in population databases in the homozygous state in asymptomatic individuals, their combination in trans results in a severe clinical phenotype. This apparent contradiction may be formally explained by intragenic complementation, a phenomenon in which two different mutant alleles of the same gene, each impairing protein function in a distinct manner, compensate for one another when present in homozygosity but together produce a dysfunctional protein complex when combined as compound heterozygotes. Precedents for this mechanism have been documented in other genetic disorders; for example, the OCA2 V443I variant is associated with oculocutaneous albinism only in compound heterozygosity with a second allele, despite being found in healthy homozygotes [47]. Similarly, a synonymous ECHS1 variant (p.P163P) has been shown to cause disease exclusively in trans with a second pathogenic allele [48].In the context of the COG complex, which functions as an eight-subunit heterooligomer, it is plausible that the F430L and I609T variants each disrupt different aspects of COG5 folding, stability, or intersubunit interactions, and that their combined presence in trans leads to cumulative loss of complex function.
Meanwhile, the observed copper accumulation and associated mitochondrial dysfunction in COG5 deficiency suggest a potential therapeutic role for copper chelation strategies. Agents such as TTM, which effectively reduce bioavailable copper, could theoretically mitigate copper-induced oxidative stress. However, the translation of this approach requires careful consideration of its therapeutic window. Copper serves as an essential cofactor for numerous cellular enzymes, most notably cytochrome c oxidase of the mitochondrial respiratory chain. Overly aggressive chelation risks inducing a state of functional copper deficiency, potentially exacerbating respiratory inefficiency and compounding metabolic dysfunction. Thus, future clinical application would necessitate precise dosing regimens, coupled with rigorous monitoring of copper status and organ function.
Tsvetkov et al. [35] found that copper-dependent cell death—cuproptosis— depends on mitochondrial respiration. In this process, copper targets lipoylated proteins of the tricarboxylic acid cycle, triggering their aggregation and promoting the loss of ISC proteins, ultimately resulting in proteotoxic stress and cell death. The mitochondrial OXPHOS impairment observed in COG5-deficient cells exhibits striking similarities to these cuproptosis features, suggesting COG5 may normally protect against copper toxicity by regulating copper distribution. However, this apparent phenotypic convergence was not supported by changes in core cuproptosis-related proteins in our proteomic analysis, indicating that the OXPHOS defects in COG5 deficiency may arise through a mechanism distinct from canonical cuproptosis pathways.
While this study establishes the role of COG5 in copper homeostasis, several questions remain. The use of HEK293T and EBV-transformed lymphoblastoid cell lines in this study enabled powerful mechanistic dissection in loss-of-function of COG5, several limitations must be acknowledged. The high transfection efficiency and proliferative capacity of HEK293T cells as a kidney-derived cell line, they may lack tissue-specific factors present in cell types more commonly affected in COG-CDGs. Similarly, EBV-transformed B cells, while a readily available source of patient-derived cell line, exhibit altered signaling and metabolic landscapes due to the persistent presence of EBV viral oncoproteins, which may confound the interpretation of cellular phenotypes.
In summary, we have identified a regulatory relationship between COG5 and copper homeostasis, demonstrating that this interaction critically influences mitochondrial OXPHOS capacity. Our findings also position COG5 as an indirect engine of copper homeostasis, may contribute to MD-pathogenesis through copper-dependent impairment of respiratory chain function.
Materials and methods
Ethics statement
The patient and healthy controls were recruited by the Department of Pediatrics, Peking University First Hospital. The study was approved by the Ethics Committee of Peking University First Hospital (Ethics approval number: 2017–217). All participants or guardians have signed informed consent forms in this study.
Cell lines and culture conditions
Cell lines: The cell lines of HEK293T (RRID: CVCL_0063), AML-12 (RRID: CVCL_0063) and B95-8 (RRID: CVCL_1953) were obtained from Cell Bank of the Chinese Academy of Science (Shanghai, China). Cell lines mentioned above have been authenticated by short tandem repeat (STR) profiling identification. Peripheral blood-derived immortalized B lymphocytes were constructed by our laboratory.
Cell culture: HEK293T and their derived cell models (including knock out, overexpression, and site-directed mutagenesis cell model) were cultured in high glucose Dulbecco’s modified Eagle medium (DMEM, Thermo Fisher Scientific, Waltham, MA, USA, Cat# C11995500T) supplemented with 12% (v/v) calf serum (CS, Sigma-Aldrich, USA, Cat# B7447). 1% (v/v) penicillin/streptomycin (P/S, Beyotime Biotechnology, China, Cat# C0222) and 0.25 mg/mL amphotericin B (MP Biomedicals, Santa Ana, USA, Cat# A610030-0001) were added to the medium. For the overexpression and site-directed mutagenesis cell models, the culture medium was additionally supplemented with 1 μg/mL puromycin for selection. Immortalized B lymphocytes were cultured in RPMI-1640 medium (Thermo Fisher Scientific, Cat# R6504) with 10% (v/v) fetal bovine serum (FBS, Sigma-Aldrich, Cat# F8318). For AML-12 was cultured in Dulbecco’s modified Eagle medium/nutrient mixture F-12 (DMEM/Hams F-12 50/50 Mix, Corning, Cat# 10–092-CV) supplemented with 12% (v/v) CS (Sigma-Aldrich, Cat# B7447), 10 μg/mL insulin (Sigma-Aldrich, Cat# abs42019847), 5.5 μg/mL transferrin (Sigma-Aldrich, Cat# T8158), 40 μg/mL dexamethasone (Sangon, Cat# A601187-0005), and 5 μg/mL selenium (Sigma-Aldrich, Cat# 229865). The cells mentioned above were cultured in a CO_2_ incubator at 37 °C under 5% CO_2_.
Patient/Control-derived immortalized B lymphocytes construction
The construction of immortalized B lymphocytes was described previously [49]. Mononuclear cells from the fresh patient’s/controls’ peripheral blood were isolated by lymphocyte separation medium (Solarbio, China, Cat# P8610). B lymphocytes were immortalized by infection with Epstein-Barr virus generated from B95-8 cells [50] supplemented with 0.5 mg/mL phytohemagglutinin (Sigma-Aldrich, Cat# L1668) and 1 mg/mL cyclosporin A (BBI Life science, Cat# A6000352). Lymphocytes were cultured with RPMI-1640 medium supplemented with 20% (v/v) FBS for the first 10 days and subsequently cultured with RPMI-1640 medium supplemented with 10% FBS.
ACMG classification
Following comprehensive in silico and in vitro analyses, we performed a detailed interpretation of the identified variant for accurately assess its pathogenicity in accordance with the American College of Medical Genetics and Genomics (ACMG) guidelines [40].
Transfection and cell models generation
Knockout (KO) cell lines were generated using the CRISPR-Cas9 technique. Genome editing support for gRNAs-designing by E-CRISP (www.e-crisp.org/E-CRISP/designcrispr.html). gRNAs (gRNAs listed in S1 Table) were inserted into pX330 (RRID: Addgene_42230) and digested by restriction enzyme Bbsl (New England Biolabs, USA, Cat#R0539) to generate the COG5-targeted KO vectors. KO cells (KO1 and KO2) and control cells were obtained by transfecting HEK293T cells with COG5-targeted KO vector and empty control vector, respectively. KO1 and KO2 were generated by two different vectors, respectively.
Knockdown (KD) cell lines were established using the RNA interference technique. Sequences of the small interfering RNAs (siRNAs) and short hairpin RNAs (shRNAs) are provided in S1 Table. The shRNA constructs were cloned into the pLKO.1 vector for stable expression in target cells.
Recovery cell models (KO1 + COG5 and KO2 + COG5) and site-directed mutagenesis cell models (Mut 1: COG5 ^c.1290C>A^, Mut 2: COG5 ^c.1826T>C^) were generated by infection of the lentiviral particles, respectively. COG5 cDNA plasmid and COG5-mutant cDNA plasmid (NM_006348.5, c.1290C > A and c.1826T > C) were cloned into pLVX-IRES-ZsGreen1 vector digested by restriction enzyme EcoRI and AgeI (New England Biolabs, Cat#R0101, Cat#R3552). Lentiviral particles were generated by transfection with co-infection of envelope vector pMD2.G (RRID: Addgene_12259), packaging vector psPAX2 (RRID: Addgene_12260), and corresponding plasmids at a ratio of 1:1:2. Lipofectamine 3000 (Thermo Fisher Scientific, Cat# L3000015) was used for all transfection experiments described above. Recovery cell models and site-directed mutagenesis cell models were purified and maintained with culture medium containing 1 μg/mL puromycin (Sangon Biotech, China, Cat# A610593-0025).
Blue-native PAGE, SDS-PAGE, and immunoblotting
Blue-native PAGE for individual- and super-complexes was performed according to a previously described protocol [51]. The protein for individual OXPHOS complexes in the native state was lysed by solubilization buffer supplemented with 1 mM PMSF and 2% (v/v) Triton X-100 (Sigma-Aldrich, Cat# 9036-19-5). Lysate was separated by centrifugation for 20 mins at 20,000 g and the supernatant was removed. Protein quantitation was done by Pierce BCA Protein Assay Kit (BCA, Thermo Fisher Scientific, Cat# 23225). The solubilized proteins were separated by a 3.5%-16% gradient gel in Cathode buffer B (containing 0.02% anionic dye Coomassie blue G-250, Sigma-Aldrich, Cat# 115444) and Cathode buffer B/10 (containing 0.002% anionic dye Coomassie blue G-250) in two successive electrophoresis procedures, respectively. Aliquots of each sample were mixed with an undenatured loading buffer by the vortex. The process was carried out with a low voltage (50 V) for stacking gel, followed by a higher voltage (180 V) for gradient gel until the dye front ran off the bottom of the gel. For the protein for OXPHOS-super complexes detection were solubilized in lysis buffer supplemented with 1% (v/v) digitonin (Sigma-Aldrich, Cat# 300410) and 1 mM PMSF. The lysate was homogenized by pipette on ice for 20 minutes and was separated by centrifugation at 20,000 g for 20 mins later. The blue-native PAGE gel with a concentration gradient of 3%-11% for mitochondrial super complex assay. The solubilized proteins were separated by a 3%-11% gradient gel in Cathode buffer B/8 (containing 0.0025% anionic dye Coomassie blue G-250, Sigma-Aldrich, Cat# 115444) in electrophoresis procedures. The entire process was rigorously conducted at 4 °C or on ice and all electrophoresis fluids, reagents, and other materials were pre-cooled.
For SDS-PAGE, the whole cells or isolated mitochondria were extracted by RIPA lysis buffer (Cell Signaling Technology, USA, Cat# 9806) supplemented with 1 mM PMSF. The samples were allowed to homogenize on ice for 10 minutes after blending and mixing by pipette. Cell debris were separated by centrifugation for 10 minutes at 14,000 g and the supernatant was removed to a new tube. Protein abundance was quantified by BCA method using a bovine-specific albumin standard curve to normalize protein abundance. Aliquots of each sample extract were boiled in SDS loading buffer at 95 °C for 5 minutes and loaded to wells for subsequent electrophoresis. The gel electrophoresis was processed with a low voltage (80 V) for stacking gel, followed by a higher voltage (120 V) for separating gel until the dye front ran off the bottom of the gel.
For immunoblotting, dispersed proteins were electroblotted onto PVDF membrane (Bio-Rad, Hercules, USA, Cat# 1620177) in pre-cooled transfer buffer supported with buffer-filled sponge and wet filter paper. The membrane was blocked with 5% (w/v) no-fat milk at room temperature for 1 hour. The membrane was incubated with appropriate primary antibodies at 4 °C overnight. Following incubation, the blot was washed 3 times with TBST and incubated with species-specific secondary antibodies. The antibodies list is seen in S1 Table. The blots were detected based on horseradish peroxidase reaction by SuperSignal West Pico PLUS Chemiluminescent Substrate (Thermo Fisher Scientific, Cat# 34577). The signal is captured on a film developed in a dark room. The blots for super-complexes were detected by alkaline phosphatase (AP)-based colorimetric reaction. The membranes were incubated in AP buffer comtaining 0.4 mg/mL NBT (Sigma-Aldrich, Cat# N6876) and 0.2 mg/mL BCIP (BBI Life Science, A610072-0500) in AP buffer (0.1 M Tris-HCl, 0.1 M NaCl, 10 mM MgCl_2_, pH = 9.5) until color development was observed.
ETC complex activity assay
ETC complex activity assay was performed according to a previously described protocol [52]. Mitochondria were suspended in suspension buffer A (10 mM Tris-HCl, 320 mM sucrose, pH 7.4) and freeze-thawed three times in liquid nitrogen for 1 min. In briefly, complex I activity was measured using an NADH-based reaction in 25 mM potassium phosphate buffer (KPi buffer) supplemented with 100 µM NADH (Sigma-Aldrich, Cat# N8129), 3 mg/mL BSA (Sigma-Aldrich, Cat# A7030), 5 mM NaN_3_, and 50 µM Decylubiquinone (DUB, Sigma-Aldrich, Cat# D7911). Complex II activity was measured using a succinate-based reaction in 25 mM KPi buffer (pH 7.4) supplemented with 1 mg/mL BSA, 5 mM NaN_3_, 75 µM 2,6-diclorophenolindophenol (DCPIP, Sigma-Aldrich, Cat# D1878), and 50 µM DUB. Complex III activity was calculated using an oxi-Cytochrome C-based reaction in 25 mM KPi buffer (pH 7.4) supplemented with 20 µM oxi-Cytochrome C (Sigma-Aldrich, Cat# C2037), 0.025% Tween-20, 100 µM EDTA-Na_2_, 5 mM NaN_3_, and 100 µM Decylubiquinol (DBH_2_, Sigma-Aldrich, Cat# D7911). Complex IV activity was measured using a reduced Cytochrome C-based reaction in 25 mM KPi buffer (pH 7.0) supplemented with 50 µM reduced Cytochrome C.
Co-immunoprecipitaton (Co-IP) analysis
The entire process was performed following the manufacturer’s instructions (P2181S, Beyotime). Cells were pelleted by centrifugation at 500 g for 3 min, resuspended in 1 mL PBS, and centrifuged again under identical conditions. After thorough removal of the supernatant, the cell pellet was lysed in 600 μL of IP lysis buffer and incubated on ice for 30 min. The lysate was centrifugated at 14,000 g for 10 min at 4 °C. The supernatant was transferred to a pre-cooled tube, and the protein concentration was determined using a BCA assay. A 50 μL aliquot of the total protein lysate was denatured at 95 °C for 5 min and stored at -80 °C as the input sample. For immunoprecipitation, flag-conjugated magnetic beads were equilibrated with TBS through two wash steps on a magnetic rack. Washed beads were resuspended in TBS to their original volume and incubated with the lysate (20 μL beads per 500 μL lysate) under continuous rotation at 4 °C overnight. Beads were then captured magnetically, and the supernatant was discarded. Subsequent washes were performed three times with 600 μL IP lysis buffer under gentle resuspension. Bound proteins were eluted in 30 μL of 2 × protein loading buffer by heating at 95 °C for 5 min. The eluate was magnetically separated, and the supernatant was collected for subsequent analysis.
Copper related-treated methods
In the metal-treated experiments, where indicated, when cells were adherent and had morphologically spread, cells were treated with medium supplemented with 1 μM CuCl_2_ (Sigma-Aldrich, Cat# C3279) and 0.17 μM disulfiram (Sigma-Aldrich, Cat# D2950000) for 24 hours or 20 μM Tetrathiomolybdate (Sigma-Aldrich, Cat# 323446) for 4 hours. Cells were then collected and used for subsequent experiments. For N-acetylcysteine (NAC; Beyotime, Cat# ST2524) treatment, cells were incubated with 1 mM N-acetylcysteine for 24 hours. Cells were then collected and used for subsequent experiments.
Mitochondrial ROS production
The amount of mitochondrial ROS was measured with MitoSOX Red mitochondrial superoxide kit (Thermo Fisher Scientific, Cat#36008) according to the manufacturer’s instructions. Briefly, cells were collected and washed by Hank’s Balanced Salt Solution (HBSS, Thermo Fisher Scientific) and resuspended and dyed by working solution with 5 μM MitoSOX red reagent diluted by HBSS buffer. Cells were then incubated at 37 °C for 10 minutes. Finally, the fluorescence signal was evaluated by NovoCyte flow cytometry (Agilent, USA) at excitation = 510 nm and emission = 580 nm.
Cellular ATP production
The production of the whole cell was detected by ATP bioluminescent somatic cell assay kit (Thermo Fisher Scientific, Cat# A22066) following the protocol provided by the manufacturer. Briefly, cells were collected and destroyed by 0.22μm filtered ultrapure water. The ATP-releasing buffer was first added to a 96-well plate to release the background signal. Aliquots of the resuspended mixed samples were added to the plate. The signals were detected using a SpectraMax iD3 Multi-Mode Microplate Reader (Molecular Devices, China) at emission = 560 nm.
Mitochondria isolation
Mitochondria extraction by using a homogenization method as previously described [53]. Cells were harvested with pre-cooled phosphate-buffered saline (PBS) with a cell scraper centrifuged at 500 g for 3 minutes at 4 °C. After removing the supernatants, the pellets were washed twice with pre-cooled PBS and centrifuged under the same conditions. The pellets were chilled on ice for 2 minutes and resuspended with hypotonic homogenate buffer for appropriate times at 1,200 g for 3 minutes at 4 °C. The degree of cell fragmentation was detected with trypan blue dye to determine the percentage of dead cells to be 90% after homogenization with a Dounce homogenizer (Wheaton, USA, Cat# 357544). The homogenate was centrifuged to discard the cell debris and obtain purified mitochondria in the supernatant. For the last step, mitochondria were enriched by centrifugation at 15,000 g for 2 minutes at 4 °C and resuspended in buffer A at -80 °C until required or for subsequent use.
Proteomic analysis
Sample Preparation for Mass Spectrometry – A piece of sample was transferred into a PCT (Pressure Cycling Technology) tube. Add 30 μL lysis buffer with 6 M urea (Sigma-Aldrich, Cat# U1250) and 2 M thiourea (Sigma-Aldrich, Cat# T8656) to each tube and mix well. Then, add 5 μL of 0.2 M TCEP (Tris(2-carboxyethyl)phosphine, Adamas-beta, Shanghai, China, Cat # 61820E) and 2.5 μL of 0.8 M IAA (Iodoacetamide), followed by 12.5 μL of 0.1 M ABB (Ammonium Bicarbonate, GENERAL-REAGENT, Shanghai, China, Cat# 1066-33-7). Set the parameters to 45 kpsi, 30 seconds high pressure, 10 seconds atmospheric pressure, 90 pressure cycles at 30°C. Subsequently, dissolve 100 μg/vial trypsin protease (Hualishi Tech, Beijing, China, Cat # HLS TRY001C) in 200 μL 1 mM HCl (Hydrochloric Acid) or 50 mM HAC (Acetic Acid, concentration 0.5 μg/μL); dissolve 50 μg rLys-C protease (Recombinant Lysyl Endopeptidase C, Hualishi Tech, Cat # HLS LYS001C) in 100 μL 1 mM HCl or 50 mM HAC (concentration 0.5 μg/μL). Add 75 μL of 0.1 M TEAB (Triethylammonium Bicarbonate) to each tube, then add 5 μg (volume 10 μL) trypsin and 1.25 μg (volume 2.5 μL) rLys-C protease. Adjust the volume with 0.1 M ABB, ensuring the pH is 8.0. Set the parameters to 20 kpsi, 50 seconds high pressure, 10 seconds atmospheric pressure, 120 pressure cycles at 30°C. Transfer the sample to a 1.5 mL EP (Eppendorf) tube to terminate the digestion by adding 15 μL of 10% TFA (Trifluoroacetic Acid, Thermo Fisher Scientific, Cat # 85183) solution to each tube, ensuring a final TFA concentration of 1%. Check that the pH is 2.0-3.0.
Activate the desalting column with 200 μL MeOH (Methanol, Sigma-Aldrich, Cat # 34860) twice; equilibrate the desalting column with 200 μL 80% ACN (Acetonitrile, Thermo Fisher Scientific, Cat # A955-4), 0.1% TFA twice; wash the desalting column with 200 μL 2% ACN, 0.1% TFA twice; desalt with 200 μL 2% ACN, 0.1% TFA ten times after sample loading; collect the sample with 100 μL 40% ACN, 0.1% TFA twice. Concentrate the sample to dryness by centrifugation under 10 mBar at 40°C and reconstitute the sample to measure peptide content at A280. Adjust the peptide concentration to 0.2 μg/μL, with an injection volume of 2 μL, ensuring a sample loading amount of 400 ng for each DIA (Data-Independent Acquisition) collection.
Mass spectrometry data analysis - The mass spectrometry data were analyzed using the DIA-NN software (version 1.8.1) for database searching, with the match-between-run (MBR) function enabled. A spectral library comprising 20,456 reviewed proteins and contaminant proteins from the Swiss-Prot database (Homo sapiens) was utilized for identification. Carbamidomethylation of cysteine residues and N-terminal methionine was specified as a fixed modification, while methionine oxidation was set as a variable modification. Both qualitative and quantitative data were obtained through this comprehensive analysis, with a stringent false discovery rate (FDR) threshold of <1% applied as the filtering criterion.
Cellular copper content determination
The entire process was performed following the manufacturer’s instructions (S1075S, beyotime).
Cell Sample Preparation - Cells were harvested and washed twice with ice-cold PBS. Cell pellets were lysed on ice using BeyoLysis Buffer A (1 × 10⁶ cells/100 µl) and incubated for 10 min with occasional gentle mixing. Lysates were then centrifuged at 12,000 × g for 5 min at 4°C to remove insoluble debris. The clarified supernatants were collected for intracellular copper measurement.
Copper Measurement - Intracellular copper levels were quantified using a colorimetric copper assay kit according to the manufacturer’s instructions. A copper standard curve (0–30 µM) was generated in parallel. For each sample, 50 µl of cell lysate was mixed with 230 µl of freshly prepared Copper Reaction Working Solution and incubated at 37°C for 5 min. Absorbance was measured at 580 nm using a microplate reader. Copper concentrations were calculated based on the standard curve and normalized to total protein concentration.
ACO-2 enzyme activity determination
Mitochondria were collected as described above. The extracted mitochondria were lysed by lysed buffer (137 mM NaCl, 1% Triton X-100, 10% glycerin, 20 mM Tris-HCl, pH = 8.0) on ice for 10 minutes, followed by centrifugation at 18,000 × g for 10 minutes. Transfer the supernatant to a fresh tube to measure the concentration of protein using a spectrophotometer. Mitochondrial proteins were separated by separating gel and stacking gel. Add 5 × loading buffer to the sample and make the volume equalized by using distilled water. Pour the running buffer into the electroporator and run the gel with 200V for 120 minutes on ice. Add chromogenic solution and incubate in 37 °C for 120 minutes on a shaker. The shade of band indicates the activity of ACO-2. The Reagent and resource list is seen in S2 Table.
Fe-S cluster fluorescent assay
Fe-S cluster fluorescent assay was performed as described [54]. Fe–S fluorescent sensor plasmids were generated by cloning genes encoding N173 (Venus’s residues 1–173) or C155 (Venus’s residues 1550–243) fused to GRX2 through a 15 amino acid (GGGGS)3 linker into pCDNA3.4. N173- and C155-terminal Venus-GRX2 fusion constructs were mixed in 1:1 v/v ratio and co-transfected into cells. Fluorescence was quantified by flow cytometry.
Sample preparation for electron microscopy
Cells were harvested at approximately 80% confluency using trypsin digestion, which was promptly neutralized with complete culture medium. The cells were centrifuged at 2,000 rpm for 5 min to form a pellet. The pellet was then gently resuspended in 1 mL of calf serum (diluted in PBS to 50%) and recentrifuged under identical conditions to promote tight aggregation of the cells. Care was taken during a subsequent PBS wash not to disrupt the pellet. After complete supernatant removal, the intact cell pellet was fixed by immersion in 1 mL of freshly prepared 2.5% glutaraldehyde in PBS and stored at 4°C for 4 hours. The fixed samples were then transferred to the Electron Microscopy Laboratory of Zhejiang University for standard processing.
Quantification and statistical analysis
All quantitative experiments were performed with three independent replicates, and all quantitative data are represented as mean ± SEM. Heatmaps were performed using pheatmap R package (Version: 1.0.12; RRID: SCR_016418) in R language (Version: 4.2.3). The image analysis of gels using ImageJ software (RRID: SCR_003070). Graphs and comparison of means between the datasets were performed with Student’s t test, one-way ANOVA, or two-way ANOVA using Prism 10.0 (RRID: SCR_002798, GraphPad Software, San Diego, CA, USA).
Supporting information
S1 FigCOG5 deficiency impairs mitochondrial OXPHOS functions.A. Western blot analysis of the steady-state levels of COG5 in HEK293T cells featuring knocking out cells (KO1 and KO2), recovery cells (KO1 + COG5 and KO1 + COG5), and corresponding control cells (Control). Control cells were obtained by transfecting empty control vectors. β-Actin served as an internal control. B-C. Analysis of steady-state levels of OXPHOS complexes in HEK293T cells with KO2, KO2 + COG5, and paired control cells (B), quantified as shown in (C). 2% Triton X-100 solubilized cells were separated and visualized by 3.5%-16% blue-native PAGE. The OXPHOS complexes were detected by immunoblotting with antibodies against proteins as indicated: CI (Grim19), CII (SDHA), CIII_2_ (UQCRC2), CIV (MT-CO1), and CV (ATP5A). TOM40 serving as the internal loading control. D. Enzymatic activities of mitochondrial electron transfer chain complexes in KO2, KO2 + COG5, and paired control cells. E. Total cellular ATP production in HEK293T cells with KO2, KO2 + COG5, and paired control cells, normalized to protein concentration. F. Mitochondrial superoxide production in HEK293T cells with KO2, KO2 + COG5, and paired control cells. G. Representative transmission electron microscopy (TEM) image of Control, KOs, and rescue model cells. Cells were fixed in 2.5% glutaraldehyde. The mitochondrion and Golgi exhibit dramatic swelling, loss of cristae structure, and membrane rupture, compared to the normal condensed morphology seen in Control (arrow). Quantitative data from three independent experiments were analyzed and presented as the means ± SEM. ns, Statistical significance is denoted as ns, no significance, *p < 0.05, **p < 0.01, ***p < 0.001, ****p < 0.0001.(TIFF)
S2 FigATP7A overexpression rescues copper accumulation and mitochondrial OXPHOS defects in COG5-deficient cells.A. ATP7A mRNA expression levels were quantified by qRT–PCR in KO1, and KO1 + ATP7A cells. B-C. Western blot analysis of ATP7A protein expression inControl, KO1, and KO1 + ATP7A cells, quantified as shown in (C). Whole-cell lysates were probed with antibodies against ATP7A and β-Actin. β-Actin served as an internal control. D. Relative intracellular copper content was determined by Elisa in Control, KO1, KO1 + COG5, and KO1 + ATP7A. Copper content in cells was normalized to total protein concentration. E. Triton X-100-solubilized lysates from Control, KO1, and KO1 + ATP7A were resolved by blue native PAGE (3.5%–16% gradient). TOM40 was used as a loading control. Antibodies against CI (Grim19), CII (SDHA), CIII_2_, (UQCRC2), CIV (MT-CO1), and CV (ATP5A) were used to detect OXPHOS complexes.(TIFF)
S3 FigOptimization of tetrathiomolybdate (TTM) treatment conditions for rescuing mitochondrial OXPHOS defects in COG5-deficient cells.A-B. Relative cell numbers of Control (A) and KO1 (B) cells treated with the indicated concentrations of tetrathiomolybdate (TTM; 0, 1, 2, 4, 5, 10, 20 µM) for 0, 2, 4, 8, 12, and 24 h. Data were normalized to untreated Control cells (0 µM) and are presented as mean ± SD (n = 3). C. Blue native PAGE (3.5%–16%) analysis of OXPHOS complexes using Triton-solubilized lysates from Control and KO1 cells treated with the indicated TTM concentrations (0, 5, 10, 20, 40 µM) for 4 h. Immunoblotting was performed using antibodies against CI (Grim19), CII (SDHA), CIII₂ (UQCRC2), CIV (MT-CO1), and CV (ATP5A). D. Blue native PAGE (3.5%–16%) analysis of mitochondrial OXPHOS complexes using Triton-solubilized lysates from Control and KO1 cells treated with TTM (2 μM, 12 h). TOM40 served as an internal loading control.(TIFF)
S4 FigIntracellular copper homeostasis regulates OXPHOS complex expression levels.A-B. Blue-native PAGE (3.5%-16% gradient) analysis of OXPHOS complexes in Triton X-100-solubilized lysates from HEK293T cells (KO2 and paired controls). TOM40 served as loading control. Experimental conditions: (-) untreated; (+) cells treated with 20 μM tetrathiomolybdate (TTM) in culture medium for 4 hours. C. Blue-native PAGE and immunoblotting analysis of mitochondrial OXPHOS super-complexes (SC). Digitonin-solubilized protein was separated by 3%-11% blue-native PAGE and the SC was immunoblotting with anti-Grim19, anti-SDHA, anti-UQCRC2, anti-MTCO1 and anti-ATP5A antibodies (showed as CI, CII, CIII_2_, CIV and CV), respectively. C1 and C2 were healthy controls. TOM70 was used as an internal loading control. Data are presented as the means ± SEM. *p < 0.05, **p < 0.01, ***p < 0.001. ****p < 0.0001.(TIFF)
S5 FigClinical presentations and pedigree diagram of patients.A-C. Brain MRI of the patient. The yellow arrow marks the location of the lesion. D. Pedigree of the family. The black arrow denotes the proband. Squares represent males, circles represent females, and solid black shapes indicate affected individuals. E. Clinical manifestations of the patients assessed using the MDC (Mitochondrial Disease Criteria) scoring system.(TIFF)
S6 FigGenetic and evolutionary analysis of COG5 variants.A. Genetic characterization of the COG5 variants c.1290C > A (p.F430L) and c.1826T > C (p.L609T), including allele frequencies (1000 Genomes, gnomAD, ExAC), dbSNP identifiers (dbSNP IDs), global minor, and in silico pathogenicity predictions (SIFT, PROVEAN, MutationTaster). B. Cross-species conservation analysis of the affected amino acid residues (F430 and L609).(TIFF)
S7 FigVariant spectrum of COG5.The variant profile of COG5 in reported patients was generated using ProteinPaint.(TIFF)
S8 FigCog5 depletion reduces ATP7A protein expression in AML-12 cells.A-B. Validation of gene knockdown efficiency in AML-12 cells. mRNA levels of (A) Cog5 and (B) Atp7a were analyzed by qRT-PCR following RNAi. C-D. Western blot analysis of ATP7A protein expression inWT and Cog5-KD cells (C), quantified as shown in (D). Whole-cell lysates were probed with antibodies against ATP7A and β-Actin. β-Actin served as an internal control.(TIFF)
S9 FigThe COG complex is required for the stability of mitochondrial OXPHOS complexes.A. Validation of subunit-specific knockdown of the COG complex in HEK293 cells. mRNA levels of COG subunits (COG1–COG8) were quantified by qRT-PCR following RNAi and normalized to β-actin. B-G. Analysis of steady-state levels of OXPHOS complexes in HEK293T cells with knock down each subunit of COG complex (COG1–COG8) (B), quantified as shown in C-G. 2% Triton X-100 solubilized cells were separated and visualized by 3.5%-16% blue-native PAGE. The OXPHOS complexes were detected by immunoblotting with antibodies against proteins as indicated (CI, Grim19; CII, SDHA; CIII_2_, UQCRC2, CIV, MT-CO1; CV, ATP5A), with TOM40 serving as the internal loading control.(TIFF)
S10 FigBlue-native PAGE analysis of OXPHOS complexes in HEK293T cells and immortalized B lymphocytes.Triton X-100-solubilized lysates from (A) HEK293T cells and (B) immortalized B lymphocytes from a healthy control were resolved by blue native PAGE (3.5%–16% gradient) in the presence or absence of 500 nM tunicamycin (48-hour treatment), quantified as shown in C-D. TOM40 was used as a loading control.(TIFF)
S11 FigEffects of antioxidant treatment and copper chelation on mitochondrial oxidative stress and OXPHOS complex integrity in COG5-deficient cells.A. Mitochondrial superoxide production was measured in Control, KO1, and KO1 cells treated with or without NAC (1 mM, 24 h). Quantitative data from three independent experiments are presented as mean ± SEM. Statistical significance is indicated as ns (not significant), *p < 0.05, **p < 0.01, ***p < 0.001, ****p < 0.0001. B. Blue native PAGE (3.5%–16%) analysis of OXPHOS complexes using Triton-solubilized cell lysates from Control, KO1, KO1 + COG5 rescued cells, and KO1 cells treated with either TTM (20 μM, 4 h) or NAC (1 mM, 24 h). Antibodies against OXPHOS complexes were used for immunoblot detection, with TOM40 serving as the loading control.(TIFF)
S1 TableKey resource table.(XLSX)
S2 TableReagent and resource for ACO-2 enzyme activity determination.(DOC)
S1 DataRaw proteomics data.(XLSX)
S2 DataSummary of reported pathogenic variants in COG5-associated disorders.(XLSX)
S3 DataClinical presentation of reported COG-CDG patients.(XLSX)
S4 DataUnderlying numerical data.(XLSX)
S5 DataUncropped and unprocessed images of the gels.(PDF)
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