Effects of SLC6A8 mutation-induced creatine deficiency on cellular function in fibroblasts

Ito, Shingo; Uemura, Tatsuki; Miyano, Ayaka; Shimbo, Hiroko; Masuda, Takeshi; Goto, Tomohide; Osaka, Hitoshi; Wada, Takahito; Ohtsuki, Sumio

doi:10.1038/s41598-025-11386-z

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Published: 23 July 2025

Effects of SLC6A8 mutation-induced creatine deficiency on cellular function in fibroblasts

Shingo Ito^1,2,
Tatsuki Uemura^2,3,
Ayaka Miyano²,
Hiroko Shimbo⁴,
Takeshi Masuda^1,2,5,
Tomohide Goto⁴,
Hitoshi Osaka⁶,
Takahito Wada⁷ &
…
Sumio Ohtsuki^1,2

Scientific Reports volume 15, Article number: 26738 (2025) Cite this article

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Abstract

Creatine transporter deficiency (CTD) caused by mutations in SLC6A8 encoding the creatine transporter (CRT), leads to cerebral creatine deficiency syndromes; however, the cellular impact of CRT loss remains unclear. In this study, we investigated the consequences of the G561R mutation by examining fibroblasts using proteomics and functional assays. We observed severe intracellular creatine deficiency (> 90% reduction), leading to impaired energy metabolism (low ATP and high ADP/ATP). Proteomic analysis revealed significant alterations in the mitochondrial and extracellular vesicle pathways. Our investigation revealed impaired mitochondrial oxidative phosphorylation, reduced spare respiratory capacity, elevated oxidative stress, and significant alterations in amino acid transporter activity. Protein misfolding associated with G561R exacerbated these deficits compared to the deletion model. These findings elucidate the key pathological mechanisms induced by the CRT-G561R mutation—including energy metabolic reprogramming, mitochondrial dysfunction, and cellular stress—which significantly contribute to our understanding of the pathogenesis of creatine transporter deficiency and suggest potential therapeutic targets.

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Introduction

The solute carrier family 6 member 8 (SLC6A8) gene encodes the creatine transporter (CRT) protein, which is essential for the active transport of creatine across plasma membranes¹. Studies using Slc6a8 knockout mice have shown that SLC6A8 is critical for maintaining creatine levels in energy-demanding tissues, such as the brain, skeletal muscle, and heart^2,3,4,5. In humans, SLC6A8 also regulates creatine levels in the brain, as individuals with SLC6A8 mutations have a cerebral creatine deficiency that causes intellectual disability^6,7,8,9. Creatine is essential for the rapid regeneration and delivery of ATP via the creatine phosphate shuttle¹⁰, and ATP-dependent energy metabolism is fundamental to normal cellular functions, supporting processes from ion transport to signal transduction^11,12. Thus, SLC6A8-mediated creatine transport is essential for maintaining cellular energy homeostasis and normal cellular functions.

To date, more than 80 loss-of-function mutations have been identified in the SLC6A8 gene¹³. A functional study showed that transient transfection of SLC6A8-deficient fibroblasts with SLC6A8 constructs harboring 21 missense variants resulted in reduced creatine uptake in 19 of the variants¹⁴. Notably, western blot analysis revealed an altered molecular weight in several missense variants compared to wild-type CRT, with at least 12 variants having a lower molecular weight than wild-type CRT¹⁴. CRT has two N-glycosylation sites (Asn192 and Asn197), and the loss of glycosylation reduces the molecular weight of the CRT protein and impairs its trafficking to the plasma membrane, resulting in a reduction in creatine transport into cells¹⁵.

We previously reported a patient with creatine transporter deficiency (CTD) carrying the missense mutation c.1681G > C (Gly561Arg) in exon 12 of SLC6A8⁶. The G561R mutant CRT showed significantly reduced creatine transport activity and was predominantly localized to the intracellular membrane fraction¹⁶. This mutation leads to immature N-glycosylation caused by protein misfolding during maturation, resulting in oligomer formation and altered cellular localization¹⁶. Furthermore, fibroblasts derived from a patient with G561R-mutated CRT (CRT-G561R fibroblasts) exhibited significantly lower creatine uptake than those derived from a patient with wild-type CRT (CRT-WT fibroblasts), although quantitative proteomics has shown that CRT protein expression levels in CRT-G561R fibroblasts were comparable to those in CRT-WT fibroblasts¹⁷. These findings prompted an investigation into how creatine depletion, mediated by the CRT-G561R mutation and driven by abnormal N-glycosylation, affects cellular energy metabolism and other biological processes, thereby providing further insights into the pathophysiology of CTD.

In this study, we aimed to elucidate the effect of CRT-G561R-mediated creatine depletion on cellular energy metabolism and biological processes in fibroblasts derived from patients with CCDSs with the G561R mutation using sequential window acquisition of all theoretical fragment ion spectra (SWATH)–mass spectrometry (MS)-based quantitative proteomics and functional analysis. By focusing on the G561R mutation, we sought not only to clarify how intracellular creatine depletion leads to cellular dysfunction but also to determine how similar glycosylation defects in other CRT mutations affect cellular function, ultimately broadening our understanding of CCDSs pathophysiology.

Results

Intracellular creatine deficiency and low ATP levels in CRT-G561R fibroblasts

As previously reported, quantitative proteomics showed no change in the CRT protein expression in CRT-G561R fibroblasts; however, creatine uptake in CRT-G561R fibroblasts was reduced by more than 90% compared to CRT-WT fibroblasts¹⁷. We confirmed that the intracellular creatine levels in CRT-G561R fibroblasts were 91.4% lower than those in CRT-WT fibroblasts (Fig. 1A), suggesting that the severe reduction in creatine uptake and intracellular creatine levels is due to impaired CRT transporter function, rather than decreased CRT protein expression. CRT-G561R fibroblasts exhibited significantly lower ATP levels (6.00% lower) and higher ADP levels (1.15-fold) than those of CRT-WT fibroblasts (Figs. 1B, C). The ADP/ATP ratio, a parameter of cellular energy metabolism, was significantly higher in CRT-G561R fibroblasts (1.22-fold) than that in CRT-WT fibroblasts (Fig. 1D). These findings indicated that the CRT-G561R mutation primarily disrupts creatine transport, leading to intracellular creatine deficiency and impaired cellular energy metabolism.

Subcellular proteome in CRT-G561R fibroblasts

The reduction in the growth rate of CRT-G561R fibroblasts compared to CRT-WT fibroblasts (43.7%, Fig. S1) indicates that impaired creatine transport negatively impacts cellular function. To investigate alterations in the cellular function in CRT-G561R fibroblasts, quantitative proteomic analysis was performed following subcellular fractionation into the cytosol, crude membrane, and plasma membrane. This analysis of CRT-G561R fibroblasts revealed differential expression of 92 cytosol, 208 crude membrane, and 99 plasma membrane proteins compared to CRT-WT fibroblasts (fold change > 2.0 or < 0.5; p < 0.05) (Figs. 2A–C, Tables S1–S3). Functional annotation analysis was performed to further investigate the changes in cellular function. STRING clustering analysis of the 286 proteins that were differentially expressed in at least one of the cytosolic, crude membrane, or plasma membrane fractions of CRT-G561R fibroblasts (Fig. 2D) revealed enrichment in pathways related to the mitochondrion (GO:0005739) and extracellular vesicles (GO:1903561) (Fig. S2).

Alterations of mitochondrial ATP production and transport in CRT-G561R fibroblasts

Of the 106 proteins annotated as mitochondrion (GO:0005739), 101 were detected in the crude membrane fraction, and 97 showed an approximately twofold increase in abundance in CRT-G561R fibroblasts compared to CRT-WT fibroblasts (Fig. 3A). Metascape pathway enrichment analysis revealed that CRT-G561R fibroblasts exhibited mitochondrial and metabolic alterations through multiple coordinated mechanisms (Fig. 3B). First, oxidative phosphorylation and the TCA cycle appeared to be promoted, as indicated by the enrichment of cellular respiration (GO:0045333), the citrate cycle (hsa00020), and pyruvate metabolism (hsa00620), with most associated proteins upregulated. Additionally, KEGG pathway enrichment analysis of the proteins involved in cellular respiration (GO:0045333) suggested a significant increase in oxidative phosphorylation (hsa00190), further supporting enhanced mitochondrial energy metabolism (Figs. 3C, D). Pathway enrichment analysis also highlighted mitochondrial electron transport from cytochrome c to oxygen (GO:0006123) and respiratory-chain complex I (CORUM:2904/2914). Proteins from all five electron-transport chain complexes (I–V) were upregulated in CRT-G561R fibroblasts (Fig. 3E). Notably, the most pronounced changes were observed in Complex I, with 23 subunits increasing by more than two-fold. In comparison, 2 Complex II subunits, 1 Complex III subunit, 8 Complex IV subunits, and 3 Complex V subunits showed similar increases (Fig. 3E). These results indicate a concerted upregulation of the entire oxidative-phosphorylation machinery, particularly Complex I, and suggest a higher theoretical capacity for ATP production.

Second, alternative fuel pathways, including the carboxylic acid metabolic process (GO:0019752), amino acid metabolic process (GO:0006520), amino acid catabolic process (GO:0009063), and fatty acid beta-oxidation (GO:0034639), were enhanced, reflecting compensatory metabolic flexibility beyond glycolysis. Third, mitochondrial quality control and biogenesis were upregulated: protein degradation (R-HSA-9837999), the unfolded protein response (R-HSA-9841251), protein import (R-HSA-1268020), and biogenesis (R-HSA-1592230) were all enhanced, supporting the rapid removal of damaged proteins and the expansion of healthy mitochondria. Finally, the oxidative stress response (GO:0006979), mitochondrial protein import (GO:0006839), and homotetramer formation (GO:0051289) were enriched, highlighting adaptive upregulation of antioxidant defenses, organelle trafficking, and oligomeric complex assembly to maintain mitochondrial integrity under energetic stress.

We next examined mitochondrial ATP export to the cytosol. Inner-membrane adenine nucleotide translocases (ANT1–3), which exchange matrix ATP for cytosolic ADP, were upregulated by ~ 1.8-fold (Fig. 3F), and outer-membrane voltage-dependent anion channels (VDAC1–3), which facilitate ATP/ADP flux, increased by ~ 1.4-fold (Fig. 3G). Mitochondrial creatine kinase CKMT1, which is responsible for converting ATP to phosphocreatine, was undetectable in the crude membrane fractions of both CRT-WT and CRT-G561R cells, whereas cytosolic creatine kinase B (CK-B) was present and modestly upregulated by 1.27-fold in CRT-G561R fibroblasts (Fig. 3H). We further investigated the mRNA levels of CKMT1 in CRT-G561R fibroblasts; however, the mRNA levels of CKMT1 were below the detection limit. Thus, mitochondrial ATP export is enhanced in CRT-G561R fibroblasts through upregulation of ANTs and VDACs, but occurs via a CKMT1-independent mechanism.

Together, these changes indicate an enhanced capacity for mitochondrial ATP production and export in CRT-G561R cells; however, although this increase does not directly proportionally to the potential increase in ATP production capacity suggested by the upregulation of other mitochondrial proteins.

Reduction of mitochondrial respiration in CRT-G561R fibroblasts

Oxygen consumption rate (OCR) measures the amount of oxygen consumed by cells and mitochondria per unit time, providing an assessment of cellular respiration, particularly oxidative phosphorylation in mitochondria. To investigate changes in mitochondrial function, the OCR of CRT-G561R fibroblasts was examined using flux analysis. OCR study showed that basal respiration, ATP production-linked respiration, proton leak, and maximal respiration were not significantly different between CRT-WT and CRT-G561R fibroblasts (Figs. 4A, B). In contrast, spare respiratory capacity was 47.5% lower, and non-mitochondrial oxygen consumption was 1.50-fold higher in CRT-G561R fibroblasts (Figs. 4A, B). Based on the increase in mitochondrial protein levels (Fig. 3D-G), we hypothesized that the number of mitochondria would be higher in CRT-G561R fibroblasts, which would compensate for the mitochondrial respiration in these fibroblasts. Since mitochondrial abundance in cells is reflected by mtDNA copy number, mtDNA copy number was examined by qPCR analysis. The mtDNA copy number in CRT-G561R fibroblasts was 2.16-fold higher than that in CRT-WT fibroblasts (Fig. 4C). When normalized to mitochondrial abundance in cells, OCR parameters per mitochondrion in CRT-G561R fibroblasts showed significantly lower basal respiration (50.2%), ATP production-linked respiration (50.3%), proton leak (49.7%), maximal respiration (62.7%), and spare respiratory capacity (85.7%) than in CRT-WT fibroblasts. Non-mitochondrial oxygen consumption was not significantly different between the two fibroblast types (Fig. 4D). Thus, although the number of mitochondria increased, the respiratory function per mitochondrion decreased, suggesting that mitochondrial proliferation compensated for maintaining ATP levels in CRT-G561R fibroblasts.

Extracellular acidification rate (ECAR) monitors the decrease in medium pH resulting from the release of acidic metabolites, primarily lactate, during cellular metabolism, and thus reflects glycolytic activity. CRT-G561R fibroblasts exhibited a slightly lower basal ECAR than CRT-WT cells (Figs. 4E, F). However, following oligomycin-induced inhibition of mitochondrial ATP synthesis, ECAR in CRT-G561R cells increased significantly more than in CRT-WT cells (Figs. 4E, F). Glycolytic reserve—the ability to boost glycolysis under stress—was significantly higher in CRT-G561R than CRT-WT fibroblasts (Figs. 4E, F). In contrast, lactate release over 24 h under normal culture conditions remained unchanged between the two cell types (Fig. 4G).

Altered intracellular energy metabolism and mitochondrial function in CRT-deficient fibroblasts

Our previous study showed that the CRT-G561R mutation disrupts creatine transport and leads to the accumulation of misfolded proteins in cells^16,17. To elucidate the dominant factor, we examined intracellular energy metabolism and mitochondrial function in CRT-deficient fibroblasts (referred to as CRT-Del fibroblasts hereafter), which exhibited a lower growth rate (Fig. S3), over 90% reduced creatine uptake¹⁷and lower intracellular creatine levels (Fig. 5A) compared to CRT-WT fibroblasts. CRT-Del fibroblasts had significantly lower ATP levels (9.0%), but ADP levels and the ADP/ATP ratio were not altered compared with CRT-WT fibroblasts (Figs. 5B–D). OCR measurements showed that basal respiration (29.6%) and ATP production-linked respiration (28.1%) were reduced, but proton leak and maximal respiration were not significantly different between CRT-WT and CRT-Del fibroblasts (Figs. 5E, F). Spare respiratory capacity was 1.67-fold higher, and non-mitochondrial oxygen consumption was 1.59-fold higher in CRT-Del fibroblasts (Figs. 5E, F). mtDNA copy number was 1.70-fold higher in CRT-Del fibroblasts than in CRT-WT fibroblasts (Fig. 5G). When normalized to mitochondrial abundance in cells, OCR parameters per mitochondrion showed that CRT-Del fibroblasts had significantly lower basal respiration (58.6%), ATP production-linked respiration (57.7%), and proton leak (63.6%), but maximal respiration, spare respiratory capacity, and non-mitochondrial oxygen consumption were not significantly altered compared to CRT-WT fibroblasts (Fig. 5H). CRT-Del fibroblasts exhibited a slightly lower basal ECAR than CRT-WT cells (Figs. 5I, J). However, following oligomycin-induced inhibition of mitochondrial ATP synthesis, the ECAR in CRT-Del fibroblasts increased significantly more than that of CRT-WT cells (Figs. 5I, J). Glycolytic reserve—the ability to boost glycolysis under stress—was significantly higher in CRT-Del fibroblasts than CRT-WT fibroblasts (Figs. 5I, J). In contrast, lactate release over 24 h under normal culture conditions remained unchanged between the two cell types (Fig. 5K).

These findings indicate that CRT-G561R fibroblasts primarily exhibit impaired creatine transport, while misfolded proteins also partially contribute to mitochondrial dysfunction, resulting in more severe mitochondrial and energetic deficits compared to CRT-Del fibroblasts.

Increase of oxidative stress-activated Nrf2 signaling in CRT-G561R fibroblasts

Oxidative stress is defined as excess reactive oxygen species (ROS) production. Mitochondrial respiratory chain defects lead to increased ROS production¹⁸, which in turn leads to cell signaling protein modification¹⁹. To evaluate ROS production, we analyzed our proteomic data for changes in the expression of oxidative stress-related proteins. Among the proteins related to “response to oxidative stress,” expression of the antioxidant enzymes superoxide dismutase 2 (SOD2) and catalase (CAT) had increased (Fig. 6A). SOD2 and CAT expression are regulated by nuclear factor erythroid 2-related factor 2 (Nrf2) signaling, which is activated in response to increases in ROS levels^20,21,22. We hypothesized that respiratory chain defects in CRT-G561R fibroblasts elevate ROS production, resulting in Nrf2 activation. To examine this hypothesis, we used STRING clustering analysis to identify proteins directly interacting with Nrf2. Among 286 differentially expressed proteins (Fig. 2D), 15 were found to directly interact with Nrf2 (Fig. 6B). Among these, five proteins were associated with “antioxidant activity” (GO:0016209). SOD2, NAD(P)H quinone dehydrogenase 1 (NQO1), thioredoxin reductase 1 (TXNRD1), peroxiredoxin 3 (PRDX3), and glutaredoxin (GLRX) protein levels were found to be higher in CRT-G561R fibroblasts than in CRT-WT fibroblasts (Fig. 6C). Therefore, these five proteins are regulated by oxidative stress–activated Nrf2 signaling.

sEV secretion from CRT-G561R fibroblasts

Proteome analysis suggested that proteins related to “Extracellular vesicle (GO:1903561)” were altered in CRT-G561R fibroblasts (Fig. S2). The abundance of proteins related to “Extracellular organelles” was classified as either increased or decreased (Fig. 7A). GO enrichment analysis using STRING revealed that both increased and decreased proteins were associated with small extracellular vesicles (sEVs), as “Exosomes (GO:0070062)” were enriched among the increased proteins and “Extracellular vesicles (GO:1903561)” were enriched among the decreased proteins (Fig. 7B). Among the 105 proteins related to “Extracellular vesicle (GO:1903561)” in the CM fraction, seven exosome marker proteins were identified, including annexin A5 (ANXA5), CD9, thrombospondin 1 (THBS1), alpha-2-macroglobulin (A2M), stomatin (STOM), lysosomal associated membrane protein 2 (LAMP2), and CD63, which are in the top 100 exosome marker proteins listed in ExoCarta (http://www.exocarta.org/). The levels of the seven identified exosome marker proteins were significantly higher in CRT-G561R fibroblasts than in CRT-WT fibroblasts (Fig. 7C). To assess the release of exosomes from fibroblasts, extracellular exosome levels were measured. Using enzyme-linked immunosorbent assay (ELISA) for both CD9 and CD63, extracellular exosome levels in the culture medium were significantly decreased in CRT-G561R fibroblasts compared to those in CRT-WT fibroblasts (Fig. 7D).

Alternations of SLC transporters in CRT-G561R fibroblasts

SLC transporters transport essential small molecules by indirectly using ATP or relying on electrochemical gradients maintained by ATP-dependent pumps. Quantitative proteomic analysis indicated that the levels of the amino acid transporters SLC7A5 and SLC38A2 showed the greatest decrease and increase, respectively, in CRT-G561R fibroblasts, as compared to the levels in CRT-WT fibroblasts (Fig. 8A). SLC7A5, an l-type amino acid transporter 1 (LAT1), forms a heterodimeric complex with 4F2hc (also known as SLC3A2) in the cell plasma membrane. The SLC3A2 protein expression levels in CRT-G561R fibroblasts were lower than those in CRT-WT fibroblasts (Fig. 8B). SLC7A5 is involved in the transport of large neutral amino acids such as leucine. [¹⁴C]-L-leucine uptake by CRT-G561R fibroblasts was found to be 31.6% lower than that of CRT-WT fibroblasts (Fig. 8C).

SLC38A2, which encodes sodium-coupled neutral amino acid transporter 2 (SNAT2), is involved in L-glutamine transport. At 48 h, glutamine consumption in the medium of CRT-G561R fibroblasts was 1.23-fold higher than that in the medium of CRT-WT fibroblasts (Fig. 8D). Thus, glutamine utilization was increased in CRT-G561R fibroblasts.

SLC1A5 is also involved in l-glutamine transport to the plasma membrane. The SLC1A5 variant plays a critical role in cancer metabolic reprogramming by transporting glutamine into mitochondria²³. To examine the changes in SLC1A5 variant expression in CRT-G561R fibroblasts, the relative ratio of unique tryptic-digested peptides derived from SLC1A5 in the PM and CM fractions was plotted. In the PM fraction, the fold changes in 5 peptides (1AA-202AA) and 11 peptides (203AA-541AA) of SLC1A5 were similar. In the CM fraction, the change in eight peptides of SLC1A5, that is, 203AA-541AA, was 1.4-fold higher than that of the two peptides 1AA-202AA; this increase was similar to that noted for VDACs (Fig. 8E), which is located in the outer mitochondrial membrane. These results may be consistent with increased expression of SLC1A5 variants in the mitochondria.

Discussion

This study investigated the cellular consequences of the G561R missense mutation in the creatine transporter (SLC6A8) using patient-derived fibroblasts. We found severe intracellular creatine deficiency (> 90% reduction), causing impaired energy metabolism (low ATP, high ADP/ATP). Proteomic analysis revealed significant alterations in mitochondrial and extracellular vesicle pathways. Our key findings demonstrate increased mitochondrial mass alongside reduced respiratory function per mitochondrion, decreased spare respiratory capacity, elevated oxidative stress, and altered amino acid transport. Furthermore, comparison with CRT-Del fibroblasts suggests that protein misfolding associated with the G561R mutation exacerbates these deficits.

Our findings confirmed severe intracellular creatine deficiency in CRT-G561R fibroblasts (> 90% reduction). This depletion directly correlated with impaired cellular energy, evidenced by lower ATP and a higher ADP/ATP ratio, highlighting a bioenergetic crisis in cells lacking intracellular creatine²⁴. Creatine plays a critical role in ATP buffering, particularly via the creatine phosphate shuttle. This can supply approximately 80% of mitochondrial ATP and enable rapid ATP production-linked respiration²⁵, which explains why CRT defects lead to severe dysfunction, especially in energy-intensive tissues³. The reduced growth rate of CRT-G561R fibroblasts further supports that impaired creatine transport negatively affects cellular vitality and proliferation, likely due to this energy limitation. When intracellular creatine is severely reduced, cells rely more heavily on less efficient non-creatine phosphate shuttle pathways for ATP supply, which provide only about 20% of mitochondrial ATP. This resulting energy stress activates AMPK, a key regulator of cellular energy homeostasis^26,27. AMPK activation drives metabolic adaptations that restore energy balance, including increased glucose uptake, fatty-acid oxidation, and mitochondrial biogenesis, while suppressing ATP-consuming processes¹¹. Quantitative proteomic analysis of CRT-G561R fibroblasts revealed significant cellular adaptations and dysfunctions, particularly a striking enrichment and upregulation of proteins in mitochondrial pathways. Reflecting this AMPK-mediated response to energy stress, these fibroblasts showed enhanced mitochondrial biogenesis and broad metabolic rewiring, including the upregulation of proteins involved in oxidative phosphorylation, the TCA cycle, pyruvate metabolism, and respiratory-chain components, alongside increased amino-acid and fatty-acid catabolism, collectively aimed at maximizing ATP generation under creatine-deficient conditions.

However, functional assessment of mitochondrial respiration (OCR) presented a complex scenario. While total basal and maximal respiration rates appeared similar between G561R and WT fibroblasts, normalizing these rates to significantly increased mtDNA copy number in G561R cells showed reduced respiratory function per mitochondrion. This indicates increased mitochondrial protein and mtDNA represent compensatory proliferation, not increased individual mitochondrial efficiency. The reduced respiratory function per mitochondrion suggests the increased mitochondrial mass operates near its maximal capacity, leaving little reserve for increased energy demands. The significantly reduced spare respiratory capacity in CRT-G561R fibroblasts confirms a diminished ability to meet sudden energy increases. This reduced flexibility is detrimental, particularly in tissues like the brain, with fluctuating energy needs²⁸. The enhanced glycolytic reserve in CRT-G561R fibroblasts supports a shift towards glycolysis when mitochondrial respiration is challenged, serving as another compensatory mechanism.

Mitochondrial membrane proteins, including VDACs and ANTs, facilitate ATP/ADP transport across the mitochondrial membranes²⁹. Although ANT1–3 and VDAC1–3 were upregulated at the total protein level in CRT-G561R fibroblasts, their relative abundance per mitochondrion was likely decreased due to the marked increase in mitochondrial number. These findings suggest that, despite an overall increase in ANT and VDAC proteins, their reduced abundance per mitochondrion may lead to decreased ATP release from individual mitochondria into the cytosol. Consequently, mitochondrial ATP export efficiency on a per-organelle basis is likely impaired, which could contribute to the observed cellular energy deficit in CRT-G561R fibroblasts.

Reduced respiratory function per mitochondrion in CRT-G561R fibroblasts likely causes increased ROS, triggering excessive proton gradient leading to over-reduction of upstream electron transport chain components (Complexes I/III), increasing electron leakage to oxygen and superoxide production, contributing to oxidative stress³⁰. Proteomics showed disproportionate upregulation of respiratory chain subunits: Complex I components were ~ 2.5-fold higher, whereas Complexes III, IV, and V were ~ 2.2-fold higher. This imbalance likely creates an electron flux bottleneck, further restricting spare respiratory capacity and impairing mitochondrial adaptability.

Beyond the primary defect in creatine transport, we observed significant alterations in other SLC transporters in CRT-G561R fibroblasts. Proteomic analysis revealed decreased expression of SLC7A5 (LAT1) and its partner SLC3A2 (4F2hc), which was functionally validated by reduced leucine uptake, indicating impaired large neutral amino acid transport. Concurrently, expression of SLC38A2 (SNAT2), a key L-glutamine transporter, was significantly upregulated, suggesting enhanced glutamine uptake. L-glutamine plays a critical role in cellular energy production by fueling the TCA cycle and supporting ATP synthesis^31,32. This increased glutamine uptake via SLC38A2, alongside potential increased mitochondrial variants of SLC1A5 transporting glutamine into mitochondria for anaplerosis^23,33, likely represents a metabolic adaptation. This metabolic reprogramming allows for a continuous supply of substrates to sustain the TCA cycle and mitochondrial ATP production³⁴ under creatine-deficient conditions. These findings highlight a cellular compensatory mechanism to counteract reduced ATP buffering and underscore the intricate interplay between amino acid transporters and mitochondrial function in response to creatine deficiency²⁴.

We also observed alterations in extracellular vesicle pathways. Proteomic analysis revealed changes in exosome-related proteins: while the levels of exosome marker proteins were increased in the crude membrane fraction, the actual release of exosomes into the medium was significantly decreased. Exosomes play a critical role in intercellular communication by transporting various molecules^35,36. Reduced exosomes secretion could impair cell-to-cell signaling and may also affect the removal of waste or misfolded proteins, thereby contributing to CTD pathology. Furthermore, the accumulation of misfolded CRT in G561R cells may disrupt the biogenesis or release of sEVs, including exosomes, potentially overwhelming the protein quality control and vesicular transport systems.

Protein accumulation and activation of the unfolded protein response induce endoplasmic reticulum stress, which contributes to the progression of neurodegenerative and metabolic diseases through cytotoxicity^37,38. A key distinction between CRT-G561R and CRT-Del fibroblasts is the accumulation of misfolded proteins in the former, which is absent in the latter. This proteotoxic stress likely exacerbates cellular stress and mitochondrial dysfunction, potentially contributing to the observed mitochondrial overproduction in CRT-G561R cells. In contrast, CRT-deletion cells, which lack this proteotoxic burden, appear to experience a less severe energy crisis and maintain greater metabolic flexibility. Thus, CRT-G561R fibroblasts are subject to both impaired creatine transport and disrupted proteostasis, resulting in more pronounced mitochondrial and energy deficits compared to CRT-Del fibroblasts.

The results of this study have implications for understanding the pathophysiology of CCDSs due to G561R and potentially other misfolding-related CRT deficiencies. Detailed characterization of metabolic reprogramming, mitochondrial dysfunction, oxidative stress, and altered vesicular trafficking provides a comprehensive view of cellular consequences beyond simple creatine depletion. These deficits likely contribute to patients’ neurological symptoms. Identification of these pathways suggests potential therapeutic avenues. Strategies aimed at improving mitochondrial function, reducing oxidative stress, enhancing protein handling, or modulating amino acid metabolism could potentially ameliorate cellular dysfunction in CTD.

However, this study has several limitations. Although fibroblasts are useful for mechanistic investigations, they may not fully recapitulate the complex cellular environment and metabolic demands of neurons, which are the primary cells affected in CCDSs^{2,39,40,41,42,43}. Future studies using patient iPSC-derived neurons or animal models will be necessary to validate these findings in more disease-relevant contexts. Furthermore, while we identified changes in several pathways, the precise molecular mechanisms linking CRT dysfunction to these alterations, particularly the signaling cascades involved in mitochondrial proliferation, oxidative stress response, and sEV regulation, require further investigation.

In conclusion, this study demonstrates that the G561R mutation causes severe intracellular creatine deficiency and profound cellular dysfunction in patient fibroblasts, characterized by impaired energy metabolism, mitochondrial dysfunction despite compensatory proliferation, oxidative stress, altered sEV secretion, and metabolic reprogramming involving amino acid transport. These findings highlight complex cellular adaptations in creatine transporter deficiency and suggest potential therapeutic targets for this disorder.

Materials and methods

Fibroblasts

Human fibroblasts (CRT-G561R mutation) were derived from a male patient with CCDSs who carried a missense mutation in exon 12 of SLC6A8 (c.1681G > C; p. Gly561Arg)⁶. Human fibroblasts (CRT-deletion) were derived from a male patient with CCDSs carrying a large 19 kb deletion that encompassed exons 5 to 13 from SLC6A8, as well as exons 5 to 8 from the B cell receptor-associated protein (BAP31) gene⁴⁴. A human fibroblast expressing wild-type CRT was also sourced from the Division of Neurology, Kanagawa Children’s Medical Center (Yokohama, Japan). This study was approved by the Ethics Committees of Kumamoto University and the Kanagawa Children’s Medical Center, and all experimental procedures were performed in accordance with the relevant institutional guidelines and regulations. Informed consent was obtained from all participants and/or their legal guardian(s).

Cell culture

Human fibroblasts were cultured in low-glucose Dulbecco’s modified Eagle’s medium (DMEM; Thermo Fisher Scientific, USA) containing 10% fetal bovine serum (FBS) and 1% penicillin–streptomycin at 37 °C in 95% air and 5% CO₂.

Cell growth test

Cells (1.0 × 10³ cells/well) were seeded in 96-well culture plates and incubated for 24 and 84 h at 37 °C in 95% air and 5% CO₂. After incubation, cell counting was performed using Cell Counting Kit-8 (Dojindo, Kumamoto, Japan), following the manufacturer’s protocol.

Creatine quantification by liquid chromatography–tandem mass spectrometry (LC-MS/MS)

Stock solutions of creatine monohydrate (Sigma-Aldrich, St. Louis, MO, USA) and d₃-creatine (methyl-D3; Cambridge Isotope Laboratories, Inc., Andover, MA, USA) were dissolved in 0.1% trifluoroacetic acid (TFA). Creatine monohydrate was diluted with 0.1% TFA to obtain calibration standards. These standard solutions supplemented with d₃-creatine as internal standard were diluted with 0.1% TFA to prepare calibration standards containing from 0, 0.03, 0.1, 0.3, 1.0, 3.0, 10 pmol/injection.

Fibroblasts were cultured in six-well plates at a density of 2.0 × 10⁵ cells/well for 3 days. Subsequently, the cells were collected using a scraper, and the cell pellets were stored at −80 °C until measurement. The cell pellets were suspended in 50 µL lysis buffer (10 mM HEPES (4- (2-hydroxyethyl)−1-piperazineethanesulfonic acid)-NaOH [pH 7.9], 10 mM KCl, and 1.5 mM MgCl₂) and sonicated for 30 min by using a Bioruptor II system (Sonicbio Co., Ltd., Kanagawa, Japan). The cell lysates were centrifuged at 8,800 × g for 5 min at 4 °C. Twenty microliters of each sample were mixed with 40 µL acetonitrile and maintained under shaker conditions for 20 min at 4 °C. Subsequently, the mixture was centrifuged at 17,500 × g for 5 min at 4 °C, and the supernatant was collected. For LC-MS/MS analysis, the samples were added to d₃creatine as an internal standard and diluted with 180 µL of 0.1% TFA.

Creatine concentration was measured using a triple quadrupole mass spectrometer (API5000; AB SCIEX, USA) coupled with a high-performance liquid chromatograph (Prominence; Shimadzu, Kyoto, Japan). Chromatographic separations were achieved on a Scherzo SS-C18 column (100 × 2.0 mm, 3 μm; Imtakt, Kyoto, Japan) under isocratic conditions of 15% B mobile phase (50% acetonitrile and 50% 100 mM ammonium formate) in phase A (0.3% formic acid in water) at a flow rate of 200 µL/min. The column temperature and injection volume were 40 °C and 20 µL, respectively. Creatine was quantified using the multiple reaction monitoring method, with transitions of the parent ions to product ions (Table S4). The peak area was extracted using the Skyline software (version 22); creatine was quantified as the ratio of creatine peak area to methyl-D3 creatine peak area using a standard curve. Creatine levels were normalized to the protein levels in the cells, measured using the bicinchoninic acid (BCA) assay.

Intracellular ATP and ADP levels

Intracellular ATP and ADP levels were determined using an ADP/ATP Ratio Assay Kit (Abcam, Cambridge, MA, USA) according to the manufacturer’s protocol. Briefly, 5 × 10³ cells were seeded in 96-well black plates and incubated for 24 h at 37 °C in a CO₂ incubator. Subsequently, the cells were treated with a nucleotide-releasing buffer to release ATP, following which ATP levels were measured using an ATP-monitoring enzyme with an Infinite^® M1000 system (Tecan, Mannedorf, Switzerland). ADP levels were measured after this mixture was incubated with ADP-converting enzyme. The ADP/ATP ratio was calculated according to the manufacturer’s protocol.

Quantitative proteomic analysis

Subcellular fractionated samples (cytosol, crude membrane, and plasma membrane) were prepared using a membrane protein extraction kit (BioVision, Milpitas, CA, USA), as previously reported⁴⁵. The protein concentrations of the fractions were measured using the Pierce BCA Protein Assay Kit (Thermo Fisher Scientific). The peptide samples for each fraction used for quantitative proteomic analysis were prepared by trypsin digestion performed using a phase-transfer surfactant⁴⁶, and proteomic analysis was performed using SWATH–MS on a 5600 TripleTOF instrument (SCIEX, Framingham, MA, USA) connected to a Dionex UltiMate 3000 RSLC Nano System (Thermo Fisher Scientific), as previously reported⁴⁷. Proteins were identified with the ProteinPilot software (version 4.5; SCIEX) using MS/MS data obtained with information-dependent acquisition. The confidence of identification was controlled with a false discovery rate of less than 1%. Proteins were identified and quantified using the library-free search function in DIA-NN 1.7 with the UniProt human reference proteome⁴⁸. Protein concentrations were calculated using the MaxLFQ algorithm⁴⁹ integrated into DIA-NN.

Oxygen consumption rate (OCR) analysis

The OCR was determined using a Seahorse XFe 24 Extracellular Flux Analyzer (Agilent Technologies, Palo Alto, CA, USA) with an Agilent Seahorse XF Cell Mito Stress Test Kit (Cat# 103015-100; Agilent) according to the manufacturer’s instructions. Briefly, cells were plated at a density of 5 × 10⁴ cells/well in an XFe 24 plate and incubated with culture medium for 24 h. The culture medium was replaced with Seahorse XF base medium 1 h before measurement, following which the cells were incubated for 1 h at 37 °C in a non-CO₂ incubator. The OCR and extracellular acidification rate (ECAR) were sequentially monitored after administration of oligomycin (1 µM; Sigma-Aldrich), carbonyl cyanide-4-(trifluoromethoxy)−4-(trifluoromethoxy)-phenylhydrazone (2 µM; FCCP, Sigma-Aldrich), rotenone (0.5 µM; Sigma-Aldrich), and antimycin A (0.5 µM); Sigma-Aldrich). Data were analyzed using the XF Wave software 2.4 (Seahorse Bioscience, Agilent).

Mitochondrial DNA (mtDNA) copy number analysis

DNA was extracted from the cells using the NucleoSpin Tissue kit (Takara Bio Inc., Kusatsu, Shiga, Japan), according to the manufacturer’s protocol, and DNA concentration and purity were determined by measuring the absorbance at 260 nm and 280 nm in a spectrophotometer (Eppendorf, Germany). mtDNA copy numbers were measured using quantitative polymerase chain reaction (qPCR; QuantStudio 3, Applied Biosystems, Foster City, CA, USA), performed using the Human Mitochondrial DNA (mtDNA) Monitoring Primer Set (Takara Bio Inc.) according to the manufacturer’s protocol. Human nuclear DNA was assessed using primers for solute carrier organic anion transporter family member 2B1 (SLCO2B1) and serpin family A member 1 (SERPINA1); human mtDNA was detected using primers for NADH dehydrogenase 1 (ND1) and NADH dehydrogenase 5 (ND5). The mtDNA copy number was calculated using the mtDNA Copy Number Calculation Tool (Takara Bio Inc.).

Lactate concentration

Cells (5 × 10³ cells/well) were seeded in 96-well culture plates and incubated for 24 h at 37 °C in 95% air and 5% CO₂. Next, the medium was replaced with fresh medium, and the cells were incubated for 24 h. The lactate concentration in the medium was measured using a Lactate Assay Kit (Dojindo), following the manufacturer’s protocol.

Measurement of exosome release

To quantify exosomes, the cells were cultured for 48 h at 37 °C in 95% air and 5% CO₂ in a medium containing extracellular vesicle-depleted FBS (Thermo). The supernatants obtained after centrifugation (500 × g for 10 min, followed by 12,000 × g for 30 min) were analyzed using an enzyme-linked immunosorbent assay (ELISA) performed using a PS Capture Exosome ELISA Kit (Wako, Japan) including anti-CD9 or anti-CD63 antibodies following manufacturer’s instructions. The sEV levels were normalized to the protein concentration of cells determined using the BCA assay (Thermo Fisher Scientific).

Glutamine concentration

Cells (1 × 10⁵ cells/well) were seeded in 24-well culture plates and incubated for 24 h at 37 °C in 95% air and 5% CO₂. Next, the medium was replaced with fresh medium, and the cells were incubated for 48 h. The glutamine concentration in the medium was measured using a Glutamine Assay Kit (Dojindo), following the manufacturer’s protocol. The glutamine concentration in the medium was normalized to the protein concentration of cells in each well, as determined using the BCA assay.

Statistics

Numerical data have been expressed in terms of mean ± standard deviation. Group-wise comparisons were performed using either Student’s t-test or Welch’s t-test, as appropriate, using GraphPad PRISM9 software (GraphPad Software, San Diego, CA, USA) or Microsoft Excel software (Microsoft Corp., Redmond, WA, USA). Network analysis of differentially expressed proteins was performed using Search Tool for the Retrieval of Interacting Genes/Proteins (STRING) 12 (https://string-db.org/), a database for aggregating and visualizing protein-protein interactions^50,51Metascape (https://metascape.org/gp/index.html#/main/step1), an integrated platform for elucidating the biological context of gene lists from high-throughput experiments⁵²and Kyoto Encyclopedia of Genes and Genomes (KEGG) pathway analysis (https://www.genome.jp/kegg/pathway.html) a comprehensive resource for mapping genes and proteins to pathways to explore functional significance^53,54,55.

Data availability

The datasets used and analysed during the current study available from the corresponding author on reasonable request.

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Acknowledgements

This study was supported by AMED (Grant Number 18ak0101080) and JST-CREST (Grant Number JP171024167). We appreciate the members of Sumio Ohtsuki’s laboratory for their valuable input. We also appreciate Editage for their professional English language editing services.

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Authors and Affiliations

Department of Pharmaceutical Microbiology, Faculty of Life Sciences, Kumamoto University, 5–1 Oe-honmachi, Chuo-ku, Kumamoto, 862-0973, Japan
Shingo Ito, Takeshi Masuda & Sumio Ohtsuki
Department of Pharmaceutical Microbiology, Graduate School of Pharmaceutical Sciences, Kumamoto University, 5–1 Oe-honmachi, Chuo-ku, Kumamoto, 862-0973, Japan
Shingo Ito, Tatsuki Uemura, Ayaka Miyano, Takeshi Masuda & Sumio Ohtsuki
Department of Molecular Pathophysiology, Institute for Advanced Medical Sciences, Nippon Medical School, 1–1–5 Sendagi, Bunkyo-ku, Tokyo, 113-8602, Japan
Tatsuki Uemura
Division of Neurology, Kanagawa Children’s Medical Center, 2–138–4 Mutsukawa, Minami-ku, Yokohama, 232-0066, Japan
Hiroko Shimbo & Tomohide Goto
Institute for Advanced Biosciences, Keio University, Tsuruoka, 997-0017, Yamagata, Japan
Takeshi Masuda
Department of Pediatrics, Jichi Medical School, 3311-1 Yakushiji, Shimotsuke-shi, 329-0498, Japan
Hitoshi Osaka
Department of Medical Ethics and Medical Genetics, Graduate School of Medicine, Kyoto University, Yoshida-Konoe-cho, Sakyo-ku, Kyoto, 606- 8501, Japan
Takahito Wada

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Contributions

S.I. and T.U. conceptualized the study and performed the investigations. S.I., T.U., A.M., T.M., and S.O. developed the methodology. T.W., H.S., T.G., and H.O. provided resources. S.I. and T.U. wrote the original draft of the manuscript, and S.I., T.U., and S.O. reviewed and edited it. S.I. and S.O. supervised the project. All authors read and approved the final manuscript.

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Correspondence to Shingo Ito.

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Ito, S., Uemura, T., Miyano, A. et al. Effects of SLC6A8 mutation-induced creatine deficiency on cellular function in fibroblasts. Sci Rep 15, 26738 (2025). https://doi.org/10.1038/s41598-025-11386-z

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Received: 28 February 2025
Accepted: 09 July 2025
Published: 23 July 2025
Version of record: 23 July 2025
DOI: https://doi.org/10.1038/s41598-025-11386-z