STIM1-Mitofusin2 interactions tether mitochondria and melanosome contacts that promote melanosome maturation

Domain: chemistry
Published: 2026-03-06
Editorial outcome: major_revision

Abstract
Mitochondria form contact sites with multiple organelles to coordinate diverse cellular processes. Melanosomes, lysosome-related organelles, undergo stepwise maturation to synthesize and store melanin, but how they interact with mitochondria remains unclear. Here we show that mitochondria–melanosome contacts dynamically increase during melanosome maturation and are mediated by STIM1–MFN2 interactions. Using a NanoBiT-based reporter system, MiMSBiT (Mitochondria–Melanosome contact reporter applying NanoBiT), to monitor reversible mitochondria–melanosome contacts in living cells, we demonstrate that STIM1 localizes to melanosomes and promotes their contact with mitochondrial MFN2. A transient decrease in melanosomal lumen calcium induces STIM1 clustering and enhances its association with MFN2. These contacts locally increase mitochondrial ATP availability, leading to melanosome lumen acidification via proton channel activation. This acidification facilitates PMEL fibrillation, a key step in melanosome maturation. Together, our findings reveal a mechanism by which mitochondria–melanosome contacts regulate melanosome maturation.

Introduction
Mitochondria are essential organelles in eukaryotic cells, playing critical roles in ATP synthesis and intracellular signal transduction. To fulfill these functions, mitochondria undergo dynamic cycles of fission and fusion and interact with various organelles1-4. Among these interactions, mitochondria-endoplasmic reticulum (ER) contact has been extensively studied using electron microscopy5. Previous studies have revealed that a variety of tethering factors, including MFN26, form functional scaffolds that facilitate the transport of metabolites such as calcium and lipids7-9. Similarly, mitochondria-melanosome contact has been observed, and MFN2 has been proposed as a potential mediator of this interaction, thereby contributing to the regulation of melanosome biogenesis10. However, many aspects of MFN2 involvement in the melanosome maturation process, as well as the dynamics and physiological significance of mitochondria-melanosome contact, remain unclear.

Melanosomes are lysosome-related organelles found in melanocytes and retinal pigment cells, specialized for the synthesis, storage, and transport of melanin11-14. They originate from endosomal precursors and undergo a series of maturation stages (I to IV) before facilitating melanin deposition and transport to the cell periphery15, 16. The synthesis of melanin and the maturation of melanosomes are tightly regulated by secreted hormones, including α-melanocyte-stimulating hormone (α-MSH)17, 18. Exposure to these hormones or ultraviolet (UV) light triggers signaling pathways, such as the cyclic adenosine monophosphate (cAMP)/protein kinase A (PKA) pathway, which promote melanin production and pigmentation19-21. Melanin is synthesized through an enzymatic process catalyzed by tyrosinase, tyrosinase-related protein 1 (Trp1), and dopachrome tautomerase, all of which are localized within the lumen of melanosomes and contribute to pigmentation22. Melanosomes are known to modulate the pH of their lumen during the maturation process. De-acidification is required for tyrosinase activation, whereas preceding acidification is essential for PMEL fibrillation, which provides a scaffold for melanin deposition23. Notably, protein fibrillation is promoted under acidic conditions24. The melanosome fibril protein PMEL was the first natural amyloid described in mammals 25 and forms amyloid fibrils only under the acidic conditions characteristic of the lysosome-like melanosome lumen, facilitating melanin synthesis and deposition26. However, the impact of mitochondria-melanosome contact on melanosome lumen pH and PMEL fibrillation remains poorly understood at the molecular and functional levels.

In this study, we utilized the NanoBiT system to measure reversible mitochondria-melanosome contacts in living cells and investigate their physiological role. Our findings reveal that MFN2-STIM1-mediated mitochondria-melanosome contact regulates melanosome lumen acidification, thereby promoting PMEL fibrillation and ensuring proper melanosome maturation.

Results
Assay system for quantifying reversible mitochondria-melanosome contact – MiMSBiT
Organelle contacts are dynamic and play critical physiological roles. However, due to the limitations of existing quantification methods, analyzing the physiological functions of organelle contact dynamics in living cells remains an area for development. In a previous study, we developed a tool to quantify reversible endoplasmic reticulum (ER)-mitochondria contacts in living cells using the NanoBiT system27. In this study, we adapted the same system to measure reversible contacts between mitochondria and melanosomes. To achieve this, we generated B16F10 cells stably expressing outer mitochondrial membrane (OMM)-targeting LgBiT (a large fragment of NanoBiT fused to human TOMM20 1-33aa) and melanosome-targeting SmBiT (a small fragment of NanoBiT fused to a melanosome-targeting tag, MST, derived from mouse Mreg 1–139 aa as previously described28). This system, named MiMSBiT (Mitochondria-Melanosome contact reporter applying NanoBiT), is illustrated and characterized in Fig. 1a-e. Luminescence was measured in MiMSBiT-expressing cells, and we observed significantly higher luminescence compared to cells that did not express the MiMSBiT system (Fig. 1f). A previous study has suggested that mitochondria and melanosomes exhibit increased contact during the early stages of melanosomal biogenesis10. Based on this, we hypothesized that mitochondria-melanosome contacts may also play a functional role during melanosome maturation. Using immunostaining analysis, we observed that α-MSH stimulation increased mitochondria–melanosome contacts (Fig. 1g, h). Consistently, MiMSBiT-based luminescence measurements revealed that stimulation with either α-MSH or forskolin, both of which enhance melanosome maturation, increased mitochondria–melanosome contacts (Fig. 1i). Together, these results demonstrate that the NanoBiT system is a powerful and reliable tool for quantifying mitochondria-melanosome contacts in living cells.

MFN2 as a critical tethering factor for mitochondria-melanosome contacts during melanosome maturation
Mitochondria-melanosome contacts have been reported to be mediated by MFN2 in melanocytes10 . To determine whether the α-MSH-stimulated contact between mitochondria and melanosomes is regulated by MFN2, we examined the formation of these contacts under conditions of Mfn2 knockdown, as well as knockdown of other known mitochondrial fission and fusion factors, including Drp1, Opa1, and Mfn129. Our results showed that knockdown of Mfn2 specifically suppressed α-MSH-induced mitochondria-melanosome contact, whereas knockdown of the other factors had no significant effect (Fig. 2a-d). Further investigation revealed that the reduction in α-MSH-dependent mitochondria-melanosome contact caused by Mfn2 knockdown was rescued by re-expression of wild-type (WT) MFN2, but not by re-expression of GTPase-inactive MFN2 mutants (Fig. 2e). Since MFN2 is known to localize to both mitochondria and the endoplasmic reticulum (ER)6, 30, we investigated whether the α-MSH-stimulated contacts are specifically regulated by mitochondria-localized or ER-localized MFN2. Using mitochondria-targeted and ER-targeted MFN2 constructs, we found that the α-MSH-induced mitochondria-melanosome contact suppressed by Mfn2 knockdown was restored by re-expression of mitochondria-targeted MFN2, but not by ER-targeted MFN2 (Fig. 2f). These findings demonstrate that α-MSH-stimulated mitochondria-melanosome contacts are specifically regulated by mitochondria-localized MFN2.
Next, we investigated the impact of mitochondria-melanosome contact on melanosome pigmentation. Prolonged α-MSH stimulation of B16F10 cells promotes melanin synthesis and pigmentation through melanosome maturation. Mfn2 knockdown significantly inhibited pigmentation induced by α-MSH stimulation, and this inhibition was rescued by the re-expression of wild-type MFN2 but not by GTPase-inactive MFN2 mutant (Fig. 2g, h). Melanin synthesis is mediated by tyrosinase, an enzyme localized within melanosomes (Supplementary Fig. 1a-c). To assess the effect of MFN2 on tyrosinase activity, we used bafilomycin A1, a H+-ATPase inhibitor, which can acutely induce melanosome de-acidification and activate tyrosinase, thereby stimulating acute melanin synthesis (Supplementary Fig. 1b). Our results showed that suppression of Mfn2 did not alter tyrosinase expression levels (Fig. 2i-k) or the melanin synthesis induced by the acute activation of tyrosinase with bafilomycin A1 (Supplementary Fig. 1d, e). These findings demonstrate that MFN2 activation positively regulates mitochondria-melanosome contact as well as α-MSH-induced pigmentation, while not affecting the expression or activity of tyrosinase.

MFN2 is required for melanosome acidification and PMEL fibrillation
The acidification of the melanosome lumen at precise stages is critical for melanosome maturation. Given that MFN2 does not regulate tyrosinase expression or activity, we hypothesized that it may instead influence melanosome acidification during α-MSH-induced maturation. To test this, we used MELOPS, a melanosome-specific pH indicator31, to measure the pH of the melanosome lumen. Since bafilomycin A1 treatment, which induces melanosome de-acidification via inhibition of V(H⁺)-ATPase, increases MELOPS fluorescence, we confirmed that MELOPS accurately reflects changes in melanosome lumen pH (Supplementary Fig. 2a, b). Using this indicator, we monitored temporal changes in melanosome pH following α-MSH stimulation and observed that melanosome acidification peaked 3 hours after stimulation with α-MSH or forskolin (Fig. 3a, b and Supplementary Fig. 2a, b). To determine whether this α-MSH–dependent acidification was mediated by V(H⁺)-ATPase activity, bafilomycin A1 was added after 3 hours of α-MSH stimulation. Bafilomycin A1 treatment resulted in a marked reduction in MELOPS fluorescence. These results indicate that α-MSH–induced melanosome acidification is largely dependent on V(H⁺)-ATPase activity. Notably, the peak in melanosome acidification coincided with the highest luminescence detected by the MiMSBiT system, observed 3 hours after α-MSH stimulation (Fig. 3c and Supplementary Fig. 2c). Depletion of Mfn2 inhibited the α-MSH-dependent acidification of the melanosome lumen, and this inhibition was rescued by re-expression of wild-type MFN2 but not by GTPase-inactive MFN2 mutants (Fig. 3a, b and Supplementary Fig. 2d). To further investigate the role of MFN2 in melanosome maturation, we examined its effect on melanosome acidification and PMEL fibrillation. During melanosome maturation, PMEL undergoes cleavage to form fibrils that serve as a scaffold for melanin deposition (Fig. 3d). It is well established that the fibrillation of PMEL is promoted by the acidification of the melanosome lumen. Using PMEL fibril-specific antibodies, we observed that fibrillation peaked around 3 hours after α-MSH stimulation, coinciding with the peak in acidification and mitochondria-melanosome contact (Fig. 3e, Supplementary Fig. 2e, f). Furthermore, inhibition of Mfn2 suppressed PMEL fibrillation following 3 hours of α-MSH stimulation (Fig. 3f). These findings suggest that MFN2 promotes mitochondria-melanosome contact, thereby facilitating melanosome acidification and enhancing PMEL fibrillation during maturation.

STIM1 binds to MFN2 and functions as a tethering factor for mitochondria-melanosome contacts
Fig. 2f clearly shows that mitochondria-localized MFN2 is crucial for mitochondria-melanosome contact. However, the tethering factor on the melanosome side remains unclear. To identify the melanosome-localized tethering factor interacting with MFN2, we utilized the TurboID proximal biotin labeling approach32. Given the propensity of TurboID for non-specific labeling, the identified proteins were considered potential interactors. By conjugating TurboID to MFN2, we analyzed proteins potentially in close proximity to MFN2 under α-MSH stimulation for 3 hours, a time point when mitochondria-melanosome contact is maximally increased (Fig. 4a-d). This approach facilitated the identification of proteins biotinylated by TurboID in close proximity through quantitative proteomics based on mass spectrometry (Fig. 4d, Supplementary Fig. 3a-c, Supplementary Data 1). Among these potential interactors, we focused on STIM1, a protein previously known not to localize to mitochondria (Fig. 4d, Supplementary Fig. 3a-c). While STIM1 is widely recognized as an endoplasmic reticulum (ER)-localized protein, our immunostaining analysis revealed that it co-localizes with TRP1, a melanosome marker, in B16F10 cells (Fig. 4e, f). Further immunoprecipitation analysis showed that STIM1 transiently binds to MFN2 at 3 hours after α-MSH stimulation but dissociates by 12 hours (Fig. 4g). Additionally, knockdown of Stim1 significantly reduced α-MSH-dependent mitochondria-melanosome contacts (Fig. 4h). These results suggest that melanosome-localized STIM1 interacts with mitochondrial MFN2 to promote mitochondria-melanosome contacts during melanosome maturation.

STIM1 binds to MFN2 in response to calcium depletion and is essential for melanosome acidification and PMEL fibrillation
Since STIM1 has been shown to bind to MFN2 and regulate mitochondria-melanosome contact, we analyzed the role of STIM1 in melanosome maturation alongside MFN2. Similar to the effects observed for MFN2 suppression, Stim1 knockdown inhibited α-MSH-stimulated melanosome acidification, melanogenesis, and PMEL fibrillation (Fig. 5a-d). To further evaluate the effects of STIM1 and MFN2 on melanosome maturation and mitochondria–melanosome contact, we performed electron microscopy analysis. Following 3 hours of α-MSH stimulation, the number of fibril-containing melanosomes was increased in control cells, whereas this increase was significantly suppressed by knockdown of MFN2 or STIM1 (Fig. 5e, f). Consistently, the number of mitochondria–melanosome contacts was also reduced in Mfn2 or Stim1 knockdown cells (Supplementary Fig. 3d). While STIM1 clearly binds to MFN2 in an α-MSH-dependent manner, the mechanism regulating this interaction remains unclear. STIM1 contains a calcium-binding domain and functions as a calcium sensor. When calcium levels decrease, calcium dissociation from STIM1 calcium-binding domain induces STIM1 clustering. We hypothesized that STIM1 binds to MFN2 through clustering triggered by α-MSH stimulation. To test this, we used BN-PAGE to analyze whether MFN2-STIM1 forms a protein complex and found that MFN2 associates with STIM1 in a large complex in an α-MSH-dependent manner (Fig. 5g). Furthermore, treatment with the calcium chelator BAPTA-AM enhanced MFN2-STIM1 binding and increased mitochondria-melanosome contact (Fig. 5h, i). To further investigate the relationship between calcium levels and contact formation, we targeted the calcium indicator RCaMP to the melanosome lumen (Supplementary Fig. 3e). Using melanosome-targeted RCaMP under α-MSH stimulation, we observed a significant decrease in melanosomal calcium concentration 30 minutes after stimulation (Fig. 5j, k). These findings demonstrate that STIM1 facilitates MFN2 binding by clustering in response to calcium depletion in the melanosome lumen, thereby promoting mitochondria-melanosome contact during melanosome maturation.

Mitochondrial ATP production facilitates melanosome acidification and PMEL fibrillation
While STIM1-MFN2 binding increases mitochondria-melanosome contacts in an α-MSH-dependent manner, the physiological consequences of these contacts remained to be determined. As an initial analysis, we found that melanosome pigmentation was suppressed when mitochondrial ATP synthesis was inhibited using oligomycin, a mitochondrial Complex V inhibitor (Fig. 6a, b). Further analysis showed that oligomycin treatment suppressed α-MSH-dependent melanosome acidification and PMEL fibrillation (Fig. 6c-e) without altering TRP1 or tyrosinase expression levels (Fig. 6f-h). Moreover, pigmentation induced by bafilomycin A1 was not suppressed by oligomycin treatment, indicating that bafilomycin A1–induced pigmentation is not dependent on mitochondrial ATP production (Fig. 6i, j). These findings demonstrate that mitochondrial ATP production is crucial for facilitating melanosome acidification and PMEL fibrillation during maturation.
Consistent with these results, mitochondrial ATP production is essential for pigmentation. Acidification of the melanosome lumen is regulated by V(H⁺)-ATPase, an ATP-driven proton pump localized on the melanosome membrane33. Based on this, we hypothesized that mitochondria supply ATP to melanosomes through direct contact, transiently activating V(H⁺)-ATPase and promoting acidification. To investigate how ATP concentration around melanosomes changes during contact with mitochondria, we targeted the ATP-sensing fluorescent protein iATPSnFR234 to the cytoplasmic side of the melanosome membrane (Supplementary Fig. 3f). Cellular ATP production occurs via two main pathways: glycolysis in the cytoplasm and oxidative phosphorylation (OXPHOS) in mitochondria. To determine which pathway contributes to ATP levels around melanosomes, we analyzed the fluorescence intensity of melanosome-localized iATPSnFR2 under conditions with or without treatment with 2-deoxy-D-glucose (2-DG), an inhibitor of hexokinase, the rate-limiting enzyme in glycolysis (Fig. 7a–e). An α-MSH–dependent increase in iATPSnFR2 fluorescence was observed even in the absence of 2-DG treatment and was suppressed by oligomycin (Fig. 7b, c). A similar pattern of α-MSH–dependent signal increase and oligomycin sensitivity was also observed in the presence of 2-DG, indicating that the ATP increase around melanosomes upon α-MSH stimulation is largely derived from mitochondrial OXPHOS (Fig. 7d, e). Notably, neither 2-DG nor oligomycin treatment significantly affected mitochondria–melanosome contact (Fig. 7f). To confirm whether mitochondria-derived ATP concentration around melanosomes is regulated by mitochondria–melanosome contact, we performed knockdown of Mfn2 or Stim1 in the presence or absence of 2-DG and measured mitochondria-derived ATP levels around melanosomes. In both conditions, depletion of Mfn2 or Stim1 significantly reduced the α-MSH–induced increase in mitochondria-derived ATP concentration around melanosomes (Fig. 7g-j). These findings suggest that melanosomes establish contact with mitochondria in response to α-MSH stimulation. At MFN2-STIM1-mediated contact sites, melanosomes are likely exposed to a locally high concentration of mitochondria-derived ATP, possibly where V(H⁺)-ATPase co-localizes, thereby promoting melanosome acidification.

MFN2–STIM1 interaction is required for melanosome maturation and pigmentation in vivo
To identify the domain of MFN2 required for its interaction with STIM1, we focused on the heptad repeat domains HR1 and HR2 of MFN2. Analysis of deletion mutants revealed that MFN2 lacking the HR2 domain, but not HR1, significantly attenuated the α-MSH–dependent binding of MFN2 to STIM1 (Fig. 8a). Consistently, treatment with a previously characterized mitofusin inhibitor35 that targets the HR2 domain of MFN2 also markedly reduced the interaction between MFN2 and STIM1 (Fig. 8b). These results indicate that the HR2 domain of MFN2 is critical for its interaction with STIM1. Furthermore, inhibition of mitofusin HR2 function significantly suppressed α-MSH–dependent acidification of the melanosome lumen (Fig. 8c, d), supporting a functional link between MFN2–STIM1 binding and melanosome maturation. To examine the physiological significance of MFN2–STIM1–mediated mitochondria–melanosome contact at the in vivo level, we analyzed pigmentation in zebrafish, in which melanization is known to be regulated by α-MSH36. Treatment of zebrafish embryos with the mitofusin inhibitor at early developmental stages resulted in a marked reduction in body pigmentation compared with control embryos (Fig. 8e, f). These findings suggest that MFN2–STIM1 interaction is required for α-MSH–dependent melanization in vivo.

Methods
All animal experiments were conducted in accordance with relevant ethical regulations and were approved by the Kanazawa University Animal Experiment Committee (protocol no. AP25-084).

Cell Culture and Transfections
B16F10 and MNT1 cells were cultured in Dulbecco's Modified Eagle’s Medium (DMEM) with high glucose, supplemented with 10% fetal bovine serum (FBS). Cells were transfected with plasmids using FuGENE HD Transfection Reagent (Promega) or with siRNAs using Lipofectamine RNAiMAX Transfection Reagent (Invitrogen), according to the manufacturer's instructions.

Small Interfering RNA
Knockdown of endogenous mouse Drp1, Opa1, Mfn1, Mfn2, tyrosinase, and Stim1 was performed using the siRNAs that were transfected with Lipofectamin RNAiMAX Transfection Reagent. Nontargeting siRNA (ID:1027281), siRNA targeting mouse Mfn1 (Catalog No: SI01304387), siRNA targeting mouse Opa1 (Catalog No: SI01365721), siRNA targeting mouse Drp1 (Catalog No: SI00982226), siRNA targeting mouse Stim1 #1 (Catalog No: SI02710736), siRNA targeting mouse Stim1 #2 (Catalog No:02735068), siRNA targeting mouse Tmem33 (Catalog No: SI01451366), siRNA targeting mouse Mff (Catalog No: SI00855932), and siRNA targeting mouse tyrosinase (Catalog No:02739107) were purchased from Qiagen. siRNA targeting human STIM1 (Catalog No: SI04251709) was purchased from Qiagen. Custom-synthesized siRNA targeting mouse Mfn2: sense, 5′-UCCUCAAGGUUUAUAAGAA-3′; human MFN2: 5′-AATCCCAGAGGGCAGAACTTTGTCC-3′44 were purchased from Japan Bio Services.

Generation of MFN2 knockout cells
To generate mMFN2L-knockout B16F10 cells, CRISPR–Cas9 vectors were constructed. Single-guide RNAs (sgRNAs) targeting the mouse Mfn2 gene were designed using the online tool CRISPOR (http://crispor.org). The oligonucleotide pair corresponding to the mMfn2 guide RNA (g3: 5′-ACAGGTACCCTTTGACGTCC-3′) was annealed and ligated into the PX459 vector (Addgene plasmid #48139). B16F10 cells were transfected using Lipofectamine 3000 (Invitrogen), and KO clones were selected by serial dilution.

Plasmids and Reagents
To generate stably expressing MiMSBiT system cells, an all-in-one pT4/HB-MiMS vector, including the OMM-targeting LgBiT, melanosome-targeting SmBiT, and IRES-PuroR, was constructed in the optimized Sleeping Beauty transposon vector (Addgene #108352)45. The OMM-targeting LgBiT was fused to an N-terminal human TOMM20 (1-33 aa) and a 3xFLAG-tag and was expressed under the EF-1α promoter. The melanosome-targeting SmBiT was fused to an N-terminal MST-tag (mouse Mreg 1-139 aa), as previously described28, and a V5-tag, and was expressed under the CBh promoter along with PuroR. To generate stably expressing myc-TurboID-MFN2 cells, myc-TurboID-MFN2 were subcloned into the pCX4 retroviral vector46. The localization-restricted Mfn2, mitochondrial Mfn2 (Mfn2ActA) and ER Mfn2 (Mfn2IYFFT) were constructed as previously reported43. Human MFN2 deletion mutants lacking the HR1 (Δ371–441) or HR2 (Δ693–747) domains were generated and subcloned into lentiviral expression vectors with an HA tag for stable expression. For melanosome-specific pH indicators, MELOPS, purchased from addgene (#80151)31, and GFP (Gamillus), an acid-tolerant monomeric green fluorescent protein purchased from addgene (#124837)47 subcloned into containing IRES sequence vectors to allow for coexpression of two genes from a single transcript. For melanosome-specific calcium indicators, RCaMP, purchased from addgene (#105013) fused tyrosinase N terminal, headed for melanosomal lumen, and GFP (Gamillus) fused to MST subcloned into containing IRES sequence vectors to allow for coexpression of two genes from a single transcript. For melanosome-specific ATP sensor, iATPSnFR2 (Addgene plasmid #209652) was fused to MST at its N-terminus and subcloned into the pLKO.1 vector. HaloTag–fused iATPSnFR2 was labeled with Janelia Fluor JFX554 HaloTag Ligand (Promega, HT1030)

The following reagents were used: α-MSH (Peptide Institute, 4057-v), forskolin (TGI, F0855), melanin (sigma, M0418), bafilomycin A1 (Adipogen Life Sciences, BVT-0252-C100), biotin (nacarai, 04822-04), BAPTA-AM (nacarai, 3731-24), oligomycin (cayman,11342), 2-DG (FUJIFILM, 046-06483).

Antibodies
The following primary antibodies were used: myc (Cell Signaling Technology, 2276; 1:1,000), V5 (MBL, PM003; 1:1,000), FLAG (Sigma-Aldrich, F3165; 1:1,000), TRP1 (Santa Cruz Biotechnology, sc-166857; 1:5,000 for immunoblotting, 1:500 for immunofluorescence), ATP5a (Abcam, ab14748; 1:5,000), α-tubulin (Sigma-Aldrich, T9026; 1:5,000), OPA1 (BD Biosciences, 612606; 1:5,000), mitofusin1 (BioLegend, 831101; dilution not applicable), mitofusin2 (Santa Cruz Biotechnology, sc-100560; dilution not applicable), tyrosinase (Santa Cruz Biotechnology, sc-20035; 1:1,000), streptavidin–HRP (Vector Laboratories, SA5004; 1:1,000), STIM1 (Cell Signaling Technology, 4916S; 1:1,000 for immunoblotting, 1:500 for immunofluorescence), GAPDH (Santa Cruz Biotechnology, sc-32233; 1:5,000), SDHA (Cell Signaling Technology, 5839S; 1:1,000), and TOMM20 (Proteintech, 11 802-1-AP; 1:5,000). The anti-PMEL (fibril) antibody was used at a dilution of 1:2,000. Anti-PMEL (fibril) was gifted from Prof. Michael S. Marks48.

Immunoblotting
Proteins in the sample buffer were separated by SDS-PAGE and transferred to Immobilon PVDF membranes (IPVH00010, Millipore). Blots were probed with the indicated antibodies, and protein bands on the blot were visualized using Chemi-Lumi One L (07880-70, Nacalai Tesque) or Immobilon Western Chemiluminescent HRP Substrate (WBKLS0500, Millipore). Band images were captured using a LuminoGraph I imager (ATTO). Relative band intensities were quantified using Fiji/ImageJ software.
For PMEL blotting, following previous reports methods with modifications were conducted49. Briefly, cells were lysed in PBST buffer containing 1% Triton X-100 and protease inhibitor cocktail in PBS, served as a whole fraction, and sedimented at 125, 000 g for 30 mins at 4°C. Supernatants from the initial fractionation were saved as the Triton-X-soluble fractions, and the pellets were washed once with PBST buffer before re-extraction with SDS buffer (PBST containing 1% SDS in place of Triton X-100). After sedimentation at 125, 000 g for 30 mins at 4°C, the supernatants were saved as the insoluble fraction.

Immunostaining
Cells were fixed with 4% paraformaldehyde in phosphate-buffered saline (PBS) for 15 minutes. Fixed cells were washed three times with PBS, permeabilized with 0.1% Triton X-100 in PBS for 10 minutes then washed three times with PBS and blocked with 10% FBS in PBS for 10 minutes, all at room temperature (RT). For immunostaining, cells were incubated with the indicated primary antibodies in PBS containing 5% FBS for 2 hours at RT, washed three times with PBS, and then incubated with the appropriate Alexa Fluor-conjugated secondary antibodies in PBS containing 5% FBS for 1 hour. The sample was washed three times with PBS, and mounted with Fluorescent Mounting Medium (S3023, Dako), and then analyzed using a FV3000 confocal laser scanning microscope (Olympus). Slice images were acquired every 0.2 μm and reconstructed using Fiji/ImageJ software. For mitochondria and melanosome colocalization analysis, using ImageJ, the areas of extraction between mitochondria and melanosomes in a 25 µm x 25 µm ROI per cell were calculated.

Immunoprecipitation
After cells were cross-linked 5mM DSP in PBS at 30 min, cells were lysed with NP-40 lysis buffer (1% Nonidet P40 Substitute, 10 mM Tris–HCl pH 7.4, 150 mM NaCl, 0.5 mM EDTA, 10 mM NaF) containing protease inhibitor cocktail (25955-11, Nacalai Tesque) and centrifuged at 15,000g for 10 min at 4 ◦C. The supernatant was subjected to immunoprecipitation using the indicated antibodies and Protein G Sepharose (17–0618-01, GE Healthcare). The beads were washed four times with NP-40 lysis buffer and boiled in 2× Laemmli sample buffer (125 mM Tris–HCl pH 6.8, 10% 2-mercaptoethanol, 4% SDS, 10% sucrose, 0.01% BPB).

Lentivirus production and infection
Lenti-X 293T Cells were seeded in 6-well plates and cultured for 2 days until they reached approximately 80% confluency at the time of transfection. Cells were transfected with lentiviral plasmid together with the packaging plasmid psPA X2 and the envelope plasmid VSV-G, using FuGENE HD Transfection Reagent (Promega). Transfections were performed in Opti-MEM, and cells were maintained in DMEM (10%FBS). On the day following transfection, half of the culture medium was replaced with fresh medium. Viral supernatants were collected 48 hours after transfection, filtered through a 0.22-μm filter, and used for infection. For lentiviral infection, target cells were incubated with viral supernatant diluted in culture medium.

Generation of MiMSBiT system stably expressing cells
Mouse melanoma B16F10 cells were kindly provided by Prof. Hideya Ando from Okayama University of Science. The pT4/HB-MiMS vector and pCAG-SB100X (Addgene #127909)50 were transfected into B16F10 cells at a ratio of 10:1 using Lipofectamine 3000 at a ratio of 10:1 (MiMS:SB100X DNA (µg)). After 24 h, cells were selected with 2 μg/ml puromycin for 2 days, and single-cell clones were isolated in 96-well plates using a limiting dilution method.

Luciferase Assay using MiMSBiT cells
Luciferase assays using the MiMSBiT system were performed using the Nano-Glo Live Cell Assay; Nano-Glo Live Cell Substrate (N205B) and NanoBiT Buffer (Nano-Glo LCS Dilution Buffer) (N206B) (Promega, Madison, WI, USA) according to the manufacturer's instructions. For luminescence quantification, MiMSBiT cells were seeded at 0.5 or 1.0×104 cells per well on a 96-well white cell culture microplate (Greiner bio-one), followed by transfection of expression vectors and treatment with rotenone or antimycin A. The cell culture medium was replaced with 100 μL of buffered FluoroBrite DMEM (Gibco) or Opti-MEM, containing <1% (v/v) fetal bovine serum (FBS). 25 µL of a 1:20 mixture of Nano-Glo Live Cell Substrate (N205B) and NanoBiT Buffer was added and incubated at 37 °C for 10 minutes. For kinetic luminescence measurements, including the analysis shown in Fig. 3c, Nano-Glo Endurazine (N2570) (Promega, Madison, WI, USA) was used according to the manufacturer’s instructions. Luminescence was then measured using a Synergy LX or HTX multimode reader (BioTek).

Kinetic fluorescent probe measurements
For kinetic fluorescent probe measurements, cells were transiently transfected or stably expressing the probe. Fluorescence signals were monitored using a Synergy HTX multimode reader (BioTek).

EM Imaging
MNT1 cells were fixed in 2.5% glutaraldehyde (Electron Microscopy Sciences, catalog no. 16022-P) in DMEM supplemented with 10% FBS at 37 °C in a 5% CO incubator for 1 h. After being washed with 0.1 M phosphate buffer (0.08 M disodium hydrogen phosphate, 0.02 M sodium dihydrogen phosphate dihydrate), the cells were scraped and collected with 0.2% BSA/0.1 M phosphate buffer, followed by centrifugation at 820 g. The cells were post-fixed with 1% OsO4 (Electron Microscopy Sciences, catalog no. 19150) and 1.5% potassium ferrocyanide (Kanto Chemical, catalog no. 32338-30) in a 0.05 M phosphate buffer for 30 min. After being rinsed 3 times with H2O, the cells were stained with 1% thiocarbohydrazide (Sigma-Aldrich, catalog no. 223220) for 5 min. After being rinsed with H2O 3 times, cells were stained with 1% OsO4 in H2O for 30 min. After being rinsed with H2O 2 times at room temperature and 3 times at 50 °C, cells were treated with Walton’s lead aspartate (0.635% lead nitrate (Sigma-Aldrich, catalog no. 203580), 0.4% aspartic acid (Sigma-Aldrich, catalog no. A9256), pH 5.2) at 50 °C for 20 min. After being rinsed with H2O 3 times, the sections were followed by incubations in an ascending ethanol series (10 min each in 50% on ice and 70% on ice, 10 min each in 90%, and 5 min in 95% ethanol/ H2O at room temperature), 5 min in 100 % ethanol 4 times and 10 min in butyl 2,3-epoxypropyl ether (Fujifilm Wako pure chemical corporation) twice. This was followed by infiltration of Plain Resin-butyl 2,3-epoxypropyl ether for o/n at a 1:1 dilution. After incubating with 100% Plain Resin for 2 h. Finally, the resin was cured at 70 °C for 5 days. Plain Resin (Nisshin EM, catalog no. 395) was made by mixing 5.9 g of Component A, 5.9 g of Component B, and 0.21g of Component C. Resin blocks were trimmed with a TrimTool diamond knife (Trim 45; DiATOME). 50 nm thick ultra-thin sections made with a diamond knife (Ultra 45; DiATOME) were collected on a cleaned silicon wafer strip in a Leica Ultramicrotome (UC7). These ultra-thin sections were collected on cleaned silicon wafer strips and imaged with a scanning electron microscope (JSM-IT800SHL, JEOL). Imaging was done at 1 kV accelerating voltage, 5 kV specimen voltage, 170 pA beam current, 2,560 × 1,920 frame size, 6 mm working distance, 12.8 × 9.6 µm field of view (pixel size 5 nm), and 5.33 µs dwell time, using the Scintillator Backscattered Electron Detector in Beam Deceleration mode.

BN-PAGE
Isolated crude mitochondria were solubilized in digitonin buffer (1% digitonin, 50 mM BisTris, 6 N HCl, 50 mM NaCl, 10% w/v Glycerol, 0.001% Ponceau S, pH 7.2) for 30 minutes on ice. Samples were centrifuged at 17,400 g for 30 minutes, and then supernatant was collected. Add 5% G-250 sample additive (Invitrogen) to the supernatant so that the final concentration is 0.5 %. Then, samples were separated on 3-12% Bis-Tris NativePAGE gels (Invitrogen), transferred to the PVDF membranes (Millipore), and immunoblotted with the indicated antibodies.

Melanin quantification assay
The B16F10 cells were collected and washed once with PBS. After the cells were detached with trypsin, they were centrifuged at 1,000 x g and the supernatant was removed. The cells were then solubilized in 150 µL of 1x RIPA buffer. After that, cells were sonicated (memory 30) for 20 seconds using a sonicator. The lysates (100 µL) were transferred to a 96-well plate for measuring absorbance (300 nm). The remaining cell lysate was used to measure protein concentration. The blank was prepared using the B16F10 White strain, which has lost its ability to synthesize melanin. A standard curve was prepared using melanin (M0418, Sigma-Aldrich) dissolved in 1× RIPA buffer. The melanin content was normalized protein content measured by the BCA kit (Takara).

Biotin labelling and organelle membrane fraction purified by digitonin
TurboID-MFN2 expressing cells were grown in 100-mm dishes 24 h before experiments. After stimulation of α-MSH 10 µM for 3 hours, the cells were incubated with fresh medium containing D-Biotin for 30 min at 37◦C for labelling with α-MSH. The cells were washed 3 times with PBS. To obtain organelle membrane fraction, a digitonin fractionation assay was performed as previously described51 with some modifications. Digitonin buffer was prepared by serial dilution of 10% digitonin (wako, 043-21371) in resuspension buffer (50 mM HEPES-KOH pH 7.4, 150 mM NaCl). The cells were pelleted by centrifugation at 500 g for 3 min at 4◦C and resuspended in PBS. After removing PBS by centrifugation, cell pellets were resuspended in 0.05% digitonin buffer, and then incubated for 10 min on ice. After centrifugation at 10,000 g for 5 min at 4◦C, the supernatant containing the cytosolic proteins was removed, and the pellet was washed with resuspension buffer. After centrifugation at 10,000 g for 5 min at 4◦C again, the supernatant was removed and the pellet was resuspended in 1% digitonin buffer. After incubation for 10 min at 4◦C, the supernatant containing membrane proteins was collected by centrifugation at 10,000 g for 5 min at 4◦C. The protein concentration of each collected supernatant was determined by BCA assay (Takara) and adjusted to 0.5 μg/μl. Each supernatant was rotated with 15 μl of Dynabeads MyOne Streptavidin C1 (streptavidin-coated magnetic beads, 65001, Invitrogen) at 4◦C overnight. The beads were washed three times with RIPA lysis buffer and pelleted on a magnetic rack. For blotting, the biotin-labelled proteins were eluted by incubating in the solution containing 0.1% SDS and 2-mM biotin for 10 min at 95◦C. The pull-down proteins on beads were subjected to LC–MS/MS analysis.

Sample preparation for LC-MS/MS analysis
The protein-bound beads were transferred to a new tube, and washed with 50 mM ammonium bicarbonate buffer twice. The beads were suspended in 100 µL of 50 mM ammonium bicarbonate buffer containing 10 mM dithiothreitol (DTT) and incubated at 55 ◦C for 15 minutes. After discarding the supernatant, 100 µL of 100 mM iodoacetamide (IAA) in 50 mM ammonium bicarbonate buffer was added to the beads, and the suspension was further incubated at room temperature for 30 minutes in the dark. After discarding the supernatant, the proteins on beads were digested by 0.5 µg of Trypsin/Lys-C mix (mass spec grade, Promega) in 100 µL of 50 mM ammonium bicarbonate buffer at 37 ◦C overnight with shaking. The supernatant was collected in a new tube and acidified with 10% trifluoroacetic acid to quench the reaction, then desalted by StageTip embedded EmporeTM SDB-XC disks (GL Science) according to the previously published literature52. The desalted eluate was dried in vacuo and dissolved in 2% MeCN, 0.1% formic acid in distilled water for LC-MS/MS measurement. Biotin-treated and biotin-untreated samples were prepared in triplicates, respectively.

LC-MS/MS analysis and data processing
LC-MS/MS analysis was performed on an Ultimate 3000 RSLCnano system (Thermo Fisher Scientific) coupled to a Q Exactive hybrid quadrupole-Orbitrap mass spectrometer (Thermo Fisher Scientific) equipped with a nano electrospray ionization source. The nanoLC system was equipped with a trap column (C18 PepMap 100, 0.3 × 5 mm, 5 µm, Thermo Fisher Scientific) and an analytical column (NTCC-360/75-3-125, Nikkyo Technos, Tokyo, Japan). Peptide separation was performed using 60 min gradient of water containing 0.1% formic acid (mobile phase A) and acetonitrile containing 0.1% formic acid (mobile phase B) at a flow rate of 300 nL/min. The elution was set as follows: 0–3 min, 2% B; 3–63 min, 2%–40% B; 63–65 min, 40%–95% B; 65–75 min, 95% B; 75–77 min, 95%–2% B; 77–90 min, 2% B. Measurements were performed in duplicate. The mass spectrometer was operated in data-dependent acquisition (DDA) mode. MS parameters were set as follows: spray voltage, 2.0 kV; capillary temperature, 275 °C; S-lens RF level, 50; scan type, full MS; scan range, m/z 350–1500; resolution, 70,000; polarity, positive; automatic gain control target, 3 × 106; maximum injection time, 100 msec. MS/MS parameters were set as follows: resolution, 17,500; automatic gain control target, 1 × 105; maximum injection time, 60 msec; normalized collision energy, 27; dynamic exclusion, 15 sec; loop count, 10; isolation window, 1.6 m/z; charge exclusion, unassigned, 1, 8, >8. The identification of peptides and proteins, and label-free quantification (LFQ) of the detected peptides were performed by using Proteome Discoverer (version 3.0, Thermo Fisher Scientific). The analytical parameters used for the data processing of peptides were as follows: parent mass error tolerance, 10.0 ppm; fragment mass error tolerance, 0.02 Da; search engine, sequest HT; protein database, SwissProt (Mus musculus); enzyme name, trypsin/P (full); maximum number of missed cleavages, 2; FDR confidence <0.01; dynamic modification, oxidation (methionine), phosphorylation (serine, threonine, tyrosine), acetyl (lysine), Met-loss (methionine), and Metloss+ acetyl (methionine); static modification, carbamidomethyl (cysteine). Label-free quantification parameters for the detected peptides were set as follows: precursor quantification, precursor abundance based on area; normalization mode, none; and hypothesis test, t-test (background based). Abundance ratio and p-values were calculated by Proteome Discoverer. From the biotinylated proteins, we selected the top 25 proteins with the highest ratio that met the criteria of p ≤ 0.05 and and high FDR confidence. Proteins with no quantitative values in the biotin-untreated sample group were excluded from the results. The mass spectrometry data were deposited to the ProteomeXchange Consortium via the jPOST partner repository with the dataset identifier (JPST003803; PXD063939). Among them, candidate genes that were not localized to mitochondria or the cytoplasm were further examined using siRNA screening. A volcano plot of biotinylated proteins proximally labeled by TurboID was generated using VolcanoR53.

Zebrafish maintenance and live imaging
Zebrafish were maintained in a MEITO system (MEITOSUIEN Co. Ltd., Aichi, Japan) at 28 °C under a 14/10 h light/dark cycle. Zebrafish embryos were obtained by natural mating. Zebrafish larvae were maintained in E3 medium (5 mM NaCl, 0.17 mM KCl, 0.33 mM CaCl2, 0.33 mM MgSO4) in the presence of MF18 (20 μM) or DMSO at 28 °C under a 14/10 h light/dark cycle. Live Imaging analyses were performed by Axio Zoom. V 16 stereomicroscope (Carl Zeiss, Oberkochen, Germany). The present analysis is approved by Kanazawa University Animal Experiment Committee.

Statistics & Reproducibility
Data are presented as mean ± s.e.m. and expressed as percentages or multiples relative to the values in control cells. Statistical significance was analyzed using Prism 9 software (GraphPad Software). Statistical significance was analyzed by either one-way analysis of variance followed by Bonferroni's test, as appropriate. P values are indicated as *p < 0.05; **p < 0.01; ***p < 0.001; ****p < 0.0001; n.s., not significant. No statistical method was used to predetermine sample size. No data were excluded from the analyses. The experiments were not randomized. The investigators were not blinded to allocation during experiments and outcome assessment. All experiments were independently repeated at least three times with similar results unless otherwise stated. In experiments, representative images (including immunoblotting and immunofluorescence) are shown from experiments that were independently repeated at least three times with similar results, unless otherwise stated. For electron microscopy analyses, experiments were performed once due to technical constraints. Multiple cells and organelles were independently analyzed from the same experiment, and quantitative analyses were based on these independent cellular observations.

Peer review rounds
[{"round": 1, "reviewer_comments": {"reviewer_1": "The manuscript by Shiiba et al. investigated the mechanism by which Mitochondria-Melanosome Contact promotes\nMelasome maturation. Although the importance of the contact has been reported, this work identifies MFN2-STIM1 as the\nmolecular requirement for contact formation and its role in promoting acidification. This work is potentially significant to the\nfield. However, there is overinterpretation of the data, and the overall significance is restricted to the cell culture.\nMajor concerns:\n1. The work proposed that the contact is specific for pmel17 fibril formation, which leads to melanosome maturation induced\nby a-MSH or Forksolin. However, the BFA-induced melanin generation is not affected. The authors proposed that the BFA\ninduces immature melanosomes. However, since all conditions cause equal darkening of the cell, more discussion is\nneeded to differentiate the immature and mature melanosome and their physiological relevance. What are the specific\ndefects in BFA-induced melanosome at the cellular or more relevant organism levels?\n2. As the authors mentioned, the major limitation of the current work is the sole dependence on cell culture. MFN2 KO\nzebrafish did not have any pigmentation problem (PMID: 23840650). Related to #1, what are the physiological signals\ndriving melanosome differentiation? The significance of the work is difficult to assess without data from animals.\n3. The conclusion that mitochondria provide local ATP is not supported by the data in Figure 7, which shows that a-MSH did\nnot simulate peri-melanosome ATP; mitochondrial ATP is important only when glycolysis is inhibited. This problem could be\ndue to the use of the MaLion sensor, which is not great at 37C. A better ATP sensor, such as iATPnFR2\n(doi.org/10.1073/pnas.2314604121)may provide better resolution.\n4. The manuscript utilizes numerous sensors that alter fluorescence intensity. The readout of the sensors should be\ndeltaF/F.\n5. It would be important to inhibit the H+-ATPase at 3 hr post-stimulation to validate that this is the major mechanism driving\npacification.\n6. The baseline of MiMSBiT is of some concern. Why is the TRP1 increase in size in Figure 1C upon MiMSBiT expression?\nThe baseline luminescence is about 600, and the activation increases the value to 1000. At the same time, the activation\nincreases mito-melanosome colocalization from 5 to 18 um2. Are there any background luminescences that limited the\ndynamic range of the MiMSBiT measurement? Would it be possible to include a negative control using a tag to an organelle\nthat is known not to interact with the melanosome to figure out the baseline?\n7. Results in Fig. 2 indicate that MFN2 is not required for contact without stimulation. Is the melanosome maturation entirely\ndependent on stimulation? Again, is MFN2 requirement context dependent?\n8. Can the STIM1 KD be reduced with a melanosome-localized STIM1? Since STIM1 regulates the entire cell's calcium\nhomeostasis, the relevance of STIM1 in melanosome maturation may have additional potential mechanisms that should be\nthoroughly discussed.\nMinor:\n1. The graphic abstract only contains the fibrillation process but not the melanin production. It would be helpful to have a\ncomprehensive summary of the paper, including the results of the BFA treatment.\n2. Line 99 needs a citation.\n3. Lines 199-200, the conclusion is confusing.\n4. Line 222, the results of Fig. 7e, f are under 2-DG treatment, and the authors did not mention this point in the results.\n5. Fig. 1f missing statistical analysis.\n6. Line 723, typo.\n7. Fig 5 h, i typo. RCamMP is not MST tagged.", "reviewer_2": "The interaction between mitochondria and melanosomes has previously been shown to be regulated by MFN2. However,\nthe involvement of other proteins and the specific roles of these membrane contact sites have remained unclear. This\nmanuscript identifies STIM1 as an additional regulator of mitochondria–melanosome membrane contact sites and provides\nevidence that these contacts promote melanosome maturation and organelle acidification. Overall, this is a high-quality and\ncomprehensive study.\nThe authors develop MiMSBiT, a tool based on the NanoBiT system, to measure mitochondria–melanosome membrane\ncontact sites. In this manuscript, it is used almost exclusively to assess the extent of these contacts. This approach supports\npreviously reported findings showing that MFN2 regulates mitochondria–melanosome contact sites, suggesting it is a\nreliable method. Nevertheless, it is important to demonstrate, even though it is described as reversible, whether this system\ninduces artificial tethering between mitochondria and melanosomes. When knocked down STIM1, it would strengthen the\nstudy to include an additional method for assessing the extent of the mitochondria–melanosome membrane contact sites.\nGiven the evidence that both MFN2 and STIM1 influence melanosome maturation, it would be good to know whether their\nknockdown results in an increased number of immature melanosomes.\nFor the experiments using MaLion and RCaMP, fluorescence images should be included to confirm correct localization to\nmelanosomes.\nLines 90-92 mentions forsklin treatment being used in the results shown in Fig 1g, h, but it is only included in Fig 1i.", "reviewer_3": "Shiiba et. al. present a manuscript describing a novel interaction between mitochondria and melanosomes that is key to\nmelanosome maturation. The authors identified the MFN2-STIM1 interaction as a driver of this mechanism. While the broad\narea of the manuscript is beyond my area of expertise, I am adequately positioned to review the interactomics data of this\nmanuscript.\nThe authors utilized a proximity labeling screen to explore for potential interacting partners for MFN2. For the purposes of\nthis manuscript, the TurboID experiments were well performed. TurboID and similar proximity labeling enzymes can be\nprofoundly non-specific, especially with the stable overexpression system utilized in this study. However, the authors\nexplicitly use this as an exploratory mechanism to ID a list of potential interactors, and they used a fractionation method\nbefore mass spectrometry to remove excess sources of background. From this they ultimately highlighted STIM1 as an\nMFN2 interactor. They followed up by directly confirming this interaction with immunoprecipitation experiments. It is\nguaranteed that some of the other hits from the screen are false positives, in fact some look familiar to me from my unrelated\nscreens (FLNA, MYO6, DHX9 to name a few). Fortunately, STIM1 was never highlighted as a false positive in my\nexperience. I encourage the authors to briefly mention the propensity for non-specific labeling with TurboID, and that the\nfindings in 4d are “potential interactors”, after which you subsequently confirmed the MFN2-STIM1 interaction. Figure 4d\ncould actually be cleaned up by comparing your data with CRAPome to help you remove some of these likely\ncontaminant protein IDs (https://doi.org/10.1038/nmeth.2557, https://reprint-apms.org/).\nAs a final point, I appreciate that the authors have deposited their raw MS data in the Proteome Xchange. However, most\nlabs do not have the tools or ability to analyze this data. It is suggested that the authors include an accessible version of their\nprocessed data highlighting the protein IDs of each sample from Proteome Discoverer. You can export the data as excel\nfiles, then combine them into a master excel file with tabs for each experiment upon resubmission."}, "author_response": "We sincerely thank all reviewers for their careful evaluation of our manuscript and for the\ninsightful and constructive comments. We have addressed each point raised by the reviewers\nin a point-by-point manner below. In particular, we greatly appreciate the suggestions that\nprompted us to incorporate in vivo zebrafish analyses, detailed ultrastructural (electron\nmicroscopy) examinations of melanosomes, and re-evaluation of ATP dynamics using improved\nsensors. The addition of morphological evidence substantially strengthened the robustness of\nour conclusions regarding melanosome maturation, while the in vivo and sensor-based\nanalyses improved the physiological relevance and technical rigor of the study. Together, these\nrevisions led to a marked improvement in the overall quality and clarity of the manuscript.\nWe hope that our revisions and responses adequately address the reviewers’ concerns, and\nwe thank them again for their valuable feedback.\nReviewer #1 (Remarks to the Author):\nThe manuscript by Shiiba et al. investigated the mechanism by which Mitochondria-\nMelanosome Contact promotes Melasome maturation. Although the importance of the contact\nhas been reported, this work identifies MFN2-STIM1 as the molecular requirement for contact\nformation and its role in promoting acidification. This work is potentially significant to the field.\nHowever, there is overinterpretation of the data, and the overall significance is restricted to\nthe cell culture.\nMajor concerns:\n1. The work proposed that the contact is specific for pmel17 fibril formation, which leads to\nmelanosome maturation induced by a-MSH or Forksolin. However, the BFA-induced melanin\ngeneration is not affected. The authors proposed that the BFA induces immature melanosomes.\nHowever, since all conditions cause equal darkening of the cell, more discussion is needed to\ndifferentiate the immature and mature melanosome and their physiological relevance. What\nare the specific defects in BFA-induced melanosome at the cellular or more relevant organism\nlevels?\nResponse:\nWe apologize for the insufficient explanation regarding bafilomycin A1 (BFA) treatment and\nmelanosome maturation in the original manuscript. Physiological melanosome maturation is\ndefined by two sequential processes: completion of PMEL17-dependent fibril formation and\nsubsequent melanin deposition (PMID: 30255724). PMEL17 fibrillation occurs in\npremelanosomes, corresponding to immature melanosomes (stages I–II), where PMEL17\nforms fibrillar scaffolds that later serve as a template for melanin deposition. In contrast,\nmature melanosomes (stages III–IV) are characterized by extensive melanin accumulation on\nthese fibrils, resulting in the loss of discernible internal matrix structures by electron\nmicroscopy. Only after proper enzyme localization, sustained melanin synthesis, and pigment\nfilling can melanosomes be considered fully mature. Bafilomycin A1 does not directly regulate\nthe melanosome maturation program itself. Instead, it elevates melanosomal luminal pH by\ninhibiting V(H⁺)-ATPase activity, which acutely enhances tyrosinase activity under neutral to\nweakly alkaline conditions (PMID: 11139343; PMID: 11461115). Therefore, in this study, BFA\nwas used to evaluate tyrosinase-dependent melanin synthesis under maturation-independent\nconditions, rather than to define or compare immature versus mature melanosomes. To avoid\nconfusion, we have removed the “immature melanosome” annotation from the BFA-treated\npanels in Extended Data Fig. 2f. The importance of fibril formation for melanosome maturation\nis well established. For example, loss of PMEL17, a major fibrillar component, abolishes\nintraluminal fibril structures, leading to impaired melanosome maturation and reduced melanin\naccumulation (PMID: 30561643; PMID: 16024012; PMID: 37833870). In zebrafish and other\nmodel organisms, PMEL17 deficiency results in clear pigmentation defects, further supporting\nfibril formation as a key determinant of functional melanosome maturation. As the reviewer\ncorrectly pointed out, our initial analyses lacked direct morphological evidence linking MFN2 or\nSTIM1 to melanosome maturation. To address this limitation, we have now performed electron\nmicroscopy analyses. These new data demonstrate that knockdown of MFN2 or STIM1\nsignificantly increases the number of melanosomes lacking fibrillar structures under α-MSH\nstimulation (Fig. 5e, f). Furthermore, to assess physiological relevance at the organismal level,\nwe performed zebrafish experiments using a mitofusin inhibitor. This treatment markedly\nreduced body surface pigmentation (Fig. 8e, f). Together, these findings support a role for\nMFN-associated pathways in melanosome maturation and pigmentation at both cellular and\norganismal levels.\n2. As the authors mentioned, the major limitation of the current work is the sole dependence\non cell culture. MFN2 KO zebrafish did not have any pigmentation problem (PMID: 23840650).\nRelated to #1, what are the physiological signals driving melanosome differentiation? The\nsignificance of the work is difficult to assess without data from animals.\nResponse:\nWe appreciate the reviewer’s concern regarding the limitation of cell culture–based analyses.\nIn the cited study (PMID: 23840650), the primary focus was on neuronal phenotypes, and\npigmentation defects in MFN2-deficient zebrafish were not examined in detail. We attempted\nto generate MFN2-deficient zebrafish to directly evaluate pigmentation; however, global MFN2\nloss caused severe developmental defects, preventing analysis of melanogenesis at later\nstages. Given our finding that the HR domain of MFN2 is essential for STIM1 binding (Fig. 8a),\nwe instead used a specific inhibitor targeting the mitofusin HR domain, which is required for\noligomerization (PMID: 35798717). Treatment of zebrafish embryos with this inhibitor resulted\nin a clear reduction in body pigmentation (Fig. 8e, f). Importantly, melanophore differentiation\nand pigmentation in zebrafish are known to be regulated by α-MSH signaling during\ndevelopment (PMID: 34502223; PMID: 40262875). Taken together, these results suggest that\nα-MSH–induced mitochondria–melanosome contact sites contribute to melanosome\nmaturation in vivo.\n3. The conclusion that mitochondria provide local ATP is not supported by the data in Figure 7,\nwhich shows that a-MSH did not simulate peri-melanosome ATP; mitochondrial ATP is\nimportant only when glycolysis is inhibited. This problem could be due to the use of the MaLion\nsensor, which is not great at 37C. A better ATP sensor, such as iATPnFR2\n(doi.org/10.1073/pnas.2314604121)may provide better resolution.\nResponse:\nWe thank the reviewer for recommending the iATPSnFR2 sensor. Using this improved sensor,\nwe reanalyzed ATP dynamics and found that α-MSH stimulation increased ATP levels at the\nmelanosome surface even in the absence of 2-DG, and this increase was suppressed by\noligomycin (Fig. 7b). These results indicate that mitochondria-derived ATP is locally enriched\naround melanosomes under α-MSH stimulation.\n4. The manuscript utilizes numerous sensors that alter fluorescence intensity. The readout of\nthe sensors should be deltaF/F.\nResponse:\nWe appreciate this suggestion. All sensor readouts have now been reanalyzed and presented\nas ΔF/F (Fig. 5a, b; Fig. 5k; Fig. 6c, d).\n5. It would be important to inhibit the H+-ATPase at 3 hr post-stimulation to validate that this\nis the major mechanism driving pacification.\nResponse:\nAs suggested, we inhibited V(H⁺)-ATPase by adding bafilomycin A1 3 hours after α-MSH\nstimulation. This treatment reversed the α-MSH–dependent decrease in melanosomal pH\n(Extended Data Fig. 2a, b), indicating that the majority of α-MSH–induced acidification is\nmediated by V(H⁺)-ATPase activity.\n6. The baseline of MiMSBiT is of some concern. Why is the TRP1 increase in size in Figure 1C\nupon MiMSBiT expression? The baseline luminescence is about 600, and the activation\nincreases the value to 1000. At the same time, the activation increases mito-melanosome\ncolocalization from 5 to 18 um2. Are there any background luminescences that limited the\ndynamic range of the MiMSBiT measurement? Would it be possible to include a negative control\nusing a tag to an organelle that is known not to interact with the melanosome to figure out\nthe baseline?\nResponse:\nWe thank the reviewer for the careful evaluation of Fig. 1c and for raising important concerns\nregarding the baseline luminescence of the MiMSBiT system. Regarding the TRP1 signal in Fig.\n1c, we repeated the immunoblot analysis under identical experimental conditions. In the repeat\nexperiments, no apparent difference in TRP1 band intensity or mobility was observed between\nparental B16F10 cells and MiMSBiT-expressing cells.\nWith respect to the baseline luminescence\nof MiMSBiT, we estimated the\nbackground level of the system using\nfunctional perturbations that disrupt\nmelanosomes. Upon suppression of\nPIKFYVE, melanosomes are lost and the\nmelanosome-targeted SmBiT becomes\ndiffusely distributed in the cytoplasm.\nUnder these conditions, MiMSBiT\nluminescence decreases to approximately\n100–200, which likely represents the background signal arising from non-specific NanoBiT\ncomplementation. In contrast, the basal luminescence observed in untreated cells is\napproximately 600-800, substantially higher than this background range. This indicates that\nthe baseline MiMSBiT signal reflects genuine basal proximity between mitochondria and\nmelanosomes rather than background luminescence. The elevated baseline further suggests\nthat, under steady-state conditions, mitochondria–melanosome contacts are partially\nmaintained by tethering mechanisms independent of MFN2 and STIM1, upon which MFN2–\nSTIM1–dependent, α-MSH–inducible contacts are superimposed. Consistent with this\ninterpretation, in our previous NanoBiT-based analysis of ER–mitochondria contacts,\nsuppression of multiple tethering-related genes reduced luminescence from the baseline level,\ndemonstrating that baseline NanoBiT signals can be genetically modulated and represent\nbiologically meaningful inter-organelle proximity (PMID: 39929810). Together, these\nobservations support the conclusion that MiMSBiT baseline luminescence reflects basal\nmitochondria–melanosome contacts rather than a limitation of the assay’s dynamic range.\n7. Results in Fig. 2 indicate that MFN2 is not required for contact without stimulation. Is the\nmelanosome maturation entirely dependent on stimulation? Again, is MFN2 requirement\ncontext dependent?\nResponse:\nOur data indicate that MFN2 is specifically required for α-MSH–induced mitochondria–\nmelanosome contacts, whereas basal contacts appear to be MFN2 independent. Thus, MFN2\nfunction is context dependent. This context dependence is consistent with the fact that\nmelanosome maturation is a hormonally regulated process rather than a constitutive event.\n8. Can the STIM1 KD be reduced with a melanosome-localized STIM1? Since STIM1 regulates\nthe entire cell's calcium homeostasis, the relevance of STIM1 in melanosome maturation may\nhave additional potential mechanisms that should be thoroughly discussed.\nResponse:\nWe agree that STIM1 regulates global calcium homeostasis. We attempted to rescue STIM1\nknockdown using melanosome-targeted STIM1 constructs; however, neither N- nor C-terminal\nMST tagging resulted in proper melanosomal localization, likely due to disruption of essential\ntargeting motifs, such as di-arginine motif and a lysine-rich region, within STIM1. We have\ntherefore expanded the Discussion to address potential indirect roles of STIM1 via cellular\ncalcium regulation (Lines 313–319).\nMinor:\n1. The graphic abstract only contains the fibrillation process but not the melanin production.\nIt would be helpful to have a comprehensive summary of the paper, including the results of\nthe BFA treatment.\nResponse:\nWe appreciate this comment. As discussed in our response to Major Comment #1, bafilomycin\nA1 induces melanin production by acutely enhancing tyrosinase activity through melanosomal\nde-acidification, rather than by promoting the physiological maturation process of\nmelanosomes. In contrast, our study focuses on mitochondria–melanosome contacts that\noccur upstream of melanin deposition and are required for proper PMEL-dependent fibril\nformation, which serves as the scaffold for subsequent melanin synthesis under α-MSH\nstimulation. Therefore, we believe that a model illustrating mitochondria–melanosome contact\nformation followed by physiological melanin production appropriately represents the core\nmechanism described in this study. The graphic abstract has been revised to clarify this\nsequence and to better reflect the scope and conclusions of our work.\n2. Line 99 needs a citation.\nResponse:\nA citation has been added.\n3. Lines 199-200, the conclusion is confusing.\nResponse:\nThis section has been revised for clarity.\n4. Line 222, the results of Fig. 7e, f are under 2-DG treatment, and the authors did not mention\nthis point in the results.\nResponse:\nFollowing additional experiments, the description of Fig. 7 has been substantially revised, and\nthe involvement of 2-DG treatment is now clearly stated in the Results section.\n5. Fig. 1f missing statistical analysis.\nResponse:\nStatistical analysis has now been added.\n6. Line 723, typo.\nResponse:\nThe typo has been corrected.\n7. Fig 5 h, i typo. RCamMP is not MST tagged.\nResponse:\nThe figure has been revised, and this issue has been corrected.\nReviewer #2 (Remarks to the Author):\nThe interaction between mitochondria and melanosomes has previously been shown to be\nregulated by MFN2. However, the involvement of other proteins and the specific roles of these\nmembrane contact sites have remained unclear. This manuscript identifies STIM1 as an\nadditional regulator of mitochondria–melanosome membrane contact sites and provides\nevidence that these contacts promote melanosome maturation and organelle acidification.\nOverall, this is a high-quality and comprehensive study.\nThe authors develop MiMSBiT, a tool based on the NanoBiT system, to measure mitochondria–\nmelanosome membrane contact sites. In this manuscript, it is used almost exclusively to\nassess the extent of these contacts. This approach supports previously reported findings\nshowing that MFN2 regulates mitochondria–melanosome contact sites, suggesting it is a\nreliable method. Nevertheless, it is important to demonstrate, even though it is described as\nreversible, whether this system induces artificial tethering between mitochondria and\nmelanosomes. When knocked down STIM1, it would strengthen the study to include an\nadditional method for assessing the extent of the mitochondria–melanosome membrane\ncontact sites.\nGiven the evidence that both MFN2 and STIM1 influence melanosome maturation, it would be\ngood to know whether their knockdown results in an increased number of immature\nmelanosomes.\nResponse:\nWe thank the reviewer for the constructive comment. To address this point, we performed\nelectron microscopy analyses. Knockdown of MFN2 or STIM1 significantly suppressed α-MSH–\ndependent fibril formation, resulting in an increased number of fibril-deficient, immature\nmelanosomes (Fig. 5e, f). In addition, electron microscopy allowed direct visualization of\nmitochondria–melanosome interfaces, providing an independent assessment of organelle\ncontacts that is not reliant on the MiMSBiT system.\nFor the experiments using MaLion and RCaMP, fluorescence images should be included to\nconfirm correct localization to melanosomes.\nResponse:\nWe confirmed melanosomal localization of RCaMP and the newly generated iATPSnFR2 by\nimmunostaining. Both sensors specifically colocalized with the melanosomal marker TRP1\n(Extended Data Fig. 3e, f).\nLines 90-92 mentions forsklin treatment being used in the results shown in Fig 1g, h, but it is\nonly included in Fig 1i.\nResponse:\nThis has been corrected; forskolin treatment is included only in Fig. 1i.\nReviewer #3 (Remarks to the Author):\nShiiba et. al. present a manuscript describing a novel interaction between mitochondria and\nmelanosomes that is key to melanosome maturation. The authors identified the MFN2-STIM1\ninteraction as a driver of this mechanism. While the broad area of the manuscript is beyond\nmy area of expertise, I am adequately positioned to review the interactomics data of this\nmanuscript.\nThe authors utilized a proximity labeling screen to explore for potential interacting partners for\nMFN2. For the purposes of this manuscript, the TurboID experiments were well performed.\nTurboID and similar proximity labeling enzymes can be profoundly non-specific, especially with\nthe stable overexpression system utilized in this study. However, the authors explicitly use this\nas an exploratory mechanism to ID a list of potential interactors, and they used a fractionation\nmethod before mass spectrometry to remove excess sources of background. From this they\nultimately highlighted STIM1 as an MFN2 interactor. They followed up by directly confirming\nthis interaction with immunoprecipitation experiments. It is guaranteed that some of the other\nhits from the screen are false positives, in fact some look familiar to me from my unrelated\nscreens (FLNA, MYO6, DHX9 to name a few). Fortunately, STIM1 was never highlighted as a\nfalse positive in my experience. I encourage the authors to briefly mention the propensity for\nnon-specific labeling with TurboID, and that the findings in 4d are “potential interactors”, after\nwhich you subsequently confirmed the MFN2-STIM1 interaction. Figure 4d could actually be\ncleaned up by comparing your data with CRAPome to help you remove some of these likely\ncontaminant protein IDs (https://doi.org/10.1038/nmeth.2557, https://reprint-apms.org/).\nAs a final point, I appreciate that the authors have deposited their raw MS data in the Proteome\nXchange. However, most labs do not have the tools or ability to analyze this data. It is\nsuggested that the authors include an accessible version of their processed data highlighting\nthe protein IDs of each sample from Proteome Discoverer. You can export the data as excel\nfiles, then combine them into a master excel file with tabs for each experiment upon\nresubmission.\nResponse:\nWe thank the reviewer for the constructive and expert comments on our interactomic analyses.\nAs pointed out, proximity labeling approaches such as TurboID can generate non-specific\nlabeling, particularly when used in stable overexpression systems, and candidate lists may\ntherefore include false-positive hits. In this study, we explicitly positioned the TurboID\nexperiment as an exploratory screening tool to identify potential MFN2-interacting proteins. To\nenhance the reliability of the dataset, we performed subcellular fractionation prior to mass\nspectrometry to enrich for mitochondria-associated fractions and reduce background noise.\nNevertheless, we agree that proteins which are frequently detected in unrelated proximity\nlabeling screens, may represent non-specific associations. Accordingly, following the reviewer’s\nsuggestion, we have added a brief mention of the potential for non-specific labeling with\nTurboID in the revised manuscript (Line 173-174, 178-179). Importantly, we subsequently\nvalidated the MFN2–STIM1 interaction by independent immunoprecipitation experiments,\nthereby confirming STIM1 as a bona fide interactor relevant to the biological mechanism\ndescribed in this study. Additionally, we realized that the results processed with a human\nFASTA database were inadvertently included in our initial manuscript. We have now replaced\nthem with the correct results using a mouse FASTA database, which is appropriate for B16F10\ncells. We apologize for this simple oversight. This corrected dataset is identical to the one\nalready uploaded to ProteomeXchange, and we confirmed that this correction did not\nsignificantly alter the overall candidate trends or the identification of STIM1. Regarding the use\nof CRAPome for contaminant filtering, we appreciate this valuable suggestion and investigated\nthe feasibility of using CRAPome. However, as far as we have investigated, the CRAPome\ndatabase is currently based primarily on human datasets, making it difficult to apply this tool\nto our mouse-derived datasets. We have clarified this limitation in the revised manuscript. We\nhope for your understanding regarding this technical limitation. Nevertheless, through the\naforementioned multi-layered validation experiments, we have made every effort to minimize\nthe influence of non-specific associations and ensure the reliability of our findings. Finally, we\nthank the reviewer for the suggestion to improve data accessibility. While the raw mass\nspectrometry data have been deposited in ProteomeXchange, we recognize that these datasets\ncan be difficult to access for non-specialists. Accordingly, we have included a newly organized\nExcel file listing the protein IDs identified via Proteome Discoverer. This supplementary table\nis intended to enhance data accessibility and ease of review for a broader range of researchers."}, {"round": 2, "reviewer_comments": {"reviewer_1": "The authors have addressed all of my concerns.", "reviewer_2": "In figure 5e were the cells in the electron microscopy images treated with MFN2 or STIM1 siRNA, as this is not labelled on\nthe figure or clear in the legend. Apart from this, the authors have adequately addressed the comments I raised, and I believe\nthe manuscript is suitable for publication.", "reviewer_3": "Overall, I am happy with the author response to my initial review. Based on my review of the interactomics data, I suggest\nthat this manuscript be accepted for publication."}, "author_response": ""}]

Decision letter