An Innovative Method for Evaluating Optical Properties of Liquid-Liquid Phase Separations using Laser Scanning Microscopic Spectroscopy

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Data may be preliminary. 27 October 2025 V1 Latest version Share on An Innovative Method for Evaluating Optical Properties of Liquid-Liquid Phase Separations using Laser Scanning Microscopic Spectroscopy Authors : Johbu Itoh , Natsuko Fujii , Saori Kohara , Masatoshi Ito , Naoto Suzuki , and Eiichiro Nagata [email protected] Authors Info & Affiliations https://doi.org/10.22541/au.176154681.12866872/v1 276 views 161 downloads Contents Abstract Supplementary Material Information & Authors Metrics & Citations View Options References Figures Tables Media Share Abstract Liquid-liquid phase separation (LLPS) has been shown to compartmentalize transcriptional condensates , thereby regulating gene expression. The optoDroplet method is frequently employed to elucidate the regulation of this process; however, the intricacies of the method remain poorly understood. The optoDroplet system, an optogenetics-based platform that uses light to activate phase transitions mediated by intrinsic disordered regions (IDRs) in living cells (NIH3T3 cells), was used to create condensed phases of fused sarcomas (FUS) driven by IDRs. The process of droplet formation and gelation of live NIH3T3 cells upon light irradiation was investigated through confocal laser microscopic spectroscopy. After exposure to light, the IDRs of FUS underwent a process of droplet formation at low light intensities, which was subsequently followed by gelation. The spectroscopic separation method enabled discrimination between monomers, dimers, and trimers, as well as gelation of the photolyase homology region (pHR)-FUS-mCherry-Cry2WT. Through the implementation of the optoDroplet method, we were able to discern disparities in the molecular morphology of the fluorescent probes. This approach may provide a novel method for elucidating the intricacies of intracellular aggregate formation. An Innovative Method for Evaluating Optical Properties of Liquid-Liquid Phase Separations using Laser Scanning Microscopic Spectroscopy Johbu Itoh 1 , Natsuko Fujii 1 , Saori Kohara 1 ， Masatoshi Ito 2 , Naoto Suzuki 1 , Eiichiro Nagata 1 ， Johbu Itoh 1 ; [email protected] , ORCID ID 0009-0003-3983-3165 Natsuko Fujii 1 ; [email protected] Saori Kohara 1 ; [email protected] Masatoshi Ito 2 ; [email protected] Naoto Suzuki 1 ; [email protected] Eiichiro Nagata 1 ; [email protected] , ORCID ID 0000-0001-6902-8309 1 Department of Neurology, Tokai University School of Medicine, Isehara, Kanagawa 259-1193 Japan 2 Department of Legal Medicine, St. Marianna University School of Medicine, Kawasaki, Kanagawa 216-8511, Japan Corresponding authors: Johbu Itoh, [email protected] Eiichiro Nagata, [email protected] Department of Neurology, Tokai University School of Medicine, Isehara, Kanagawa 259-1193 Japan Acknowledgments The authors thank the Life Science Support Center, Tokai University, for LSM880 Equipment Maintenance. Funding This work was supported by Conflicts of Interest The authors declare no conflicts of interest. Abstract Liquid-liquid phase separation (LLPS) has been shown to compartmentalize transcriptional condensates, thereby regulating gene expression. The optoDroplet method is frequently employed to elucidate the regulation of this process; however, the intricacies of the method remain poorly understood. The optoDroplet system, an optogenetics-based platform that uses light to activate phase transitions mediated by intrinsic disordered regions (IDRs) in living cells (NIH3T3 cells), was used to create condensed phases of fused sarcomas (FUS) driven by IDRs. The process of droplet formation and gelation of live NIH3T3 cells upon light irradiation was investigated through confocal laser microscopic spectroscopy. After exposure to light, the IDRs of FUS underwent a process of droplet formation at low light intensities, which was subsequently followed by gelation. The spectroscopic separation method enabled discrimination between monomers, dimers, and trimers, as well as gelation of the photolyase homology region (pHR)-FUS-mCherry-Cry2WT. Through the implementation of the optoDroplet method, we were able to discern disparities in the molecular morphology of the fluorescent probes. This approach may provide a novel method for elucidating the intricacies of intracellular aggregate formation. Keywords: Liquid-liquid phase separation (LLPS), optoDroplet, confocal laser microscopic spectroscopy Summary 1. The optoDroplet method was utilized to study the LLPS phenomenon. 2. The application of blue light to NIH3T3 cells resulted in the formation of cell droplets containing FUS within the cell. 3. The present study successfully differentiated monomers (liquefaction), dimers (liquid-phase formation), and gel formation using laser scanning microscopic spectroscopy. 4. The application of a prolonged dose of energy to cells resulted in the formation of a gel. 5. The results of this study demonstrated the potential of optical spectroscopic techniques to analyze LLPS using the optoDroplet method. In addition, a new mechanism contributing to cell death in neurodegenerative diseases was identified, providing valuable insights into this process. Research Highlights The LLPS phenomenon in the optoDroplet method can be discerned through the use of fluorescent molecular spectroscopy, which enabled the distinction between droplet formation and gelation. Introduction Recent studies have shown that various proteins and RNAs undergo liquid-liquid phase separation (LLPS) during self-assembly, forming droplets or gel-like structures in cells (Anderson, Kedersha, 2009.；Brangwynne, Mitchison, and Hyman, 2011.; Li, 2012.; BrnBrangwynne, 2013; Molliex et al., 2015; Nott et al., 2015; Shin and Brangwynne, 2017; Banani, Lee, Hyman,and Rosen, 2017; Dolgin, 2018.; Boeynaems, et al., 2018; Olzmann,and Carvalho. 2019; Zhe Feng, Xudong Chen, Xiandeng Wu, and Mingjie Zhang. 2019.; Peng, YU, Fang. 2024; Zhu, et al.2025; Huang, Dong, 2025.;). It has been suggested that cells regulate various biological processes, such as transcription, translation, and signal transduction, by forming and dissolving biomolecular droplets (Niu, 2020; Garcia-Cabau & Salvatella, 2021). As the importance of cell phase separation has become clearer, research on the chemical and physical properties of cell droplets, as well as the biological processes and diseases they regulate, has increased dramatically in recent years (Zhang et al., 2024; Jeon et al., 2025). However, analyzing the mechanisms and properties of droplet formation and the role of cells involved has been challenging using conventional methods that observe droplets only after applying stress or stimuli to cells (Antifeeva et al., 2022). Therefore, attempts have been made to manipulate cell droplet formation using light or low-molecular-weight compounds in real time to detect intracellular protein-protein interactions and control cell signaling molecules (Zhang et al., 2024). One of the methods we employ is LLPS to elucidate the mechanisms of cell death caused by intracellular aggregate formation in neurodegenerative diseases (Li et al., 2022). However, the regulation of this process remains unclear. In this paper, we report a detailed analysis of LLPS droplet and aggregate formation using the optoDroplet method (Sine et al., 2017), a chemical biology technique that generates artificial droplets in cells using light (Shaner, et al., 2004.; Banerjee, et al., 2007. ; Topol, Collins, Savitsky, Nemukhin. 2011.; Cranfill, et al., 2016. ; Ding. Philip D. Andrew. DeMello. 2019.; Ding, Howes, DeMello,. 2019) through confocal laser microscopic wavelength spectroscopy (Bigelow et al., 2003; Inomoto et al., 2007; Itoh & Osamura, 2007; Collins et al., 2011. ; Zhang, et al.,2013). Here, we report the results of this study. Materials and Methods Cell culture NIH 3T3 cells were cultured in 10% FBS (Atlanta Biological) in Dulbecco’s Modified Eagle Medium (DMEM, GIBCO) supplemented with penicillin, streptomycin, and GlutaMAX (Thermo) at 37 ℃ with 5% CO2 in a humidified incubator. Plasmid We used the optogenetics-based ”optoDroplet” system to control phase transitions mediated by IDRs in living NIH3T3 cells through photoexcitation. FUS with mCherry fluorescent protein (FP) and Arabidopsis thaliana (At) pHR Cry2 (pHR-FUSN-mCh-Cry2WT #101223, #223) (Addgene) was expressed in living NIH3T3 cells, and the response to light stimulation was observed. As a control vector, pHR-mCherry-Cry2 WT #101221(#221) (Addgene) was used. Cell imaging Live cells were plated on 35-mm glass-bottom dishes and grown twice overnight in normal growth medium to reach approximately 70% confluency. Imaging with a Confocal Laser Scanning Microscope All live-cell imaging was performed using a ZEISS LSM880 microscope (ZEISS, Jena, Germany) equipped with a 37°C temperature-controlled stage and a 5 % CO 2 atmosphere. Spectral analysis was conducted on cells cultured in 35-mm glass-bottom dishes. The samples were observed using a Plan-Apochromat lens (10×, numerical aperture 0.45, M27, Zeiss) and a C-Apochromat lens (63×, numerical aperture 1.2 water, M27, Zeiss). Time-lapse lambda-mode images were obtained using an argon laser (wavelength 488 nm, 0.03 mW) for Cry2 activation and a diode-pumped solid-state (DPSS) laser (wavelength 561 nm, 0.017 mW) for mCherry excitation. Imaging parameters were as follows: pixel dwell time, 1.54 μs; line averaging, 16; pinhole size, 33 μm; filter range, 499–696 nm; and beam splitter, MBS 488/561. ZEN imaging software (version 2.3 SP, Carl Zeiss Microscopy, Jena, Germany) was used for spectral unmixing and image processing. Fluorescence spectroscopy To examine opto-droplet formation in LLPS, we expressed pHR-mCherry-Cry2WT and pHR-FUS-mCherry-Cry2WT in NIH3T3 cells. The spectral profiles of the monomer, dimer, trimer, and gelation forms of the FP mCherry were acquired by spectroscopy over the wavelength range of 499–696 nm. We examined (1) the presence of spectral shifts, broadening, and deformation, (2) the occurrence of slight redshifts in the peak wavelength, e.g., from 610 to 612–615 nm, and (3) the presence of spectral broadening and bimodality (Figure 2b). Autofluorescence spectra from NIH3T3 cells, as well as mCherry spectra from NIH3T3 cells (#221) transfected with pHR-mCherry-Cry2WT as wild-type control cells. mCherry dimerization and gelation spectra were obtained from NIH3T3 cells transfected with pHR-FUS-mCherry-Cry2WT (#223). In #223 cells, spectral data were acquired from regions corresponding to non-photo-stimulated areas (monomer), photo-stimulated droplet formation (dimerization), and strongly photo-stimulated gelation (dimerization and gelation). Lambda stack images were acquired over the wavelength range of 499–696 nm with 9 nm intervals. Spectra for each mCherry state were separated and identified from cellular autofluorescence (e.g., 610 nm) based on waveform shape, peak value, and half-width parameters. Time-lapse scan observation For the time-lapse scan observation, images were captured at 30 s intervals for 20 or 40 cycles. The relative fluorescence intensities of monomeric mCherry (red), dimeric mCherry (green), and gel-phase mCherry (cyan) along the cell were calculated from 2D or 3D scanning data. To perform activation protocols with varying activation intervals, the first frame was set under light-off conditions (non-light activation), and light-on (light activation) cycles were applied from the second frame onward. Image Analysis ZEN Blue edition ver. 3.9 (ZEISS) and IMARIS ver. 6.1 (BitPlane) were used to measure and correct the total molecular concentration and the steady-state background concentration outside clusters from fluorescent images of cells. Briefly, raw images were averaged to remove noise, and droplets were detected based on peak intensity. All detected clusters in each video were manually inspected to ensure accuracy. The cell boundaries and nuclear periphery were manually identified, and only cytoplasmic clusters were analyzed. All intracellular fluorescence intensities were integrated to quantify the total amount of phase-separated material in each frame. Results The optogenetics-based ”optoDroplet” system, which utilizes photoexcitation to regulate phase transitions through IDRs in living cells, was employed in this study. The mCherry FP and the pHR of At were fused to FUS, and the resulting light-on/off responses were observed in living cells. We expressed pHR-mCherry-Cry2 WT and pHR-FUS-mCherry-Cry2WT in NIH3T3 cells. Subsequently, droplet (dimer) formation was induced using blue light. The following results present the fluorescence spectra of pHR-mCherry-Cry2WT and pHR-FUS-mCherry-Cry2WT as acquired through spectroscopy. Similar to previous reports (Taslimi et al., 2014; Lee et al., 2014), blue light-activated formation of clusters (droplet: dimer) was not observed in most cells expressing pHR-mCherry-Cry2WT (Figure 1a). Cry2 (photoreceptor) undergoes mild homodimerization and conformational changes in response to blue light. However, wild-type Cry2 (Cry2WT) has weak interaction affinity and limited self-assembly ability. Therefore, mCherry-Cry2WT showed diffuse fluorescence throughout the cell even after blue light stimulation, as observed in component-extracted images (602–620 nm) using a conventional bandpass filter (Figure 1a, yellow band). Conversely, when FUS (FUSN) was fused to the N-terminal IDR of Cry2WT (optoFUS; pHR-FUSN-mCherry-Cry2WT), blue light-activated cluster (droplet: dimer) formation increased markedly in most cells. In the bandpass filter component-extracted images, the fluorescent properties of monomers, dimers, trimers, and gelation were not recognizable; only the morphology revealed a substantial accumulation of blue light-activated clusters (Figure 1b). Configuration of the optoDroplet method The molecular design of the optoDroplet used in this study consists of an N-terminal LC domain derived from the naturally denatured protein of FUS (control: no LC domain), a C-terminal FP mCherry for observation, and an Arabidopsis-derived photoreceptor protein, Cry2, which forms homodimers in response to blue light. Cry2 proteins do not undergo phase separation in the dark; however, light stimulation causes Cry2 to dimerize and cluster through LC domain-mediated multi-molecular interactions, forming intracellular droplets. The amount of protein clustering in these droplets can be controlled by adjusting light exposure intensity and duration. When light exposure is ceased, Cry2 reverts to a monomer, making the droplets a reversible system. In NIH3T3 cells expressing pHR-mCherry-Cry, the molecular structure of pHR-mCherry-Cry2 WT remained unchanged before and after blue light stimulation, with mCherry fluorescence diffusely distributed throughout the cell and showing no detectable changes, as shown in its single spectrum (Figure 2a). In NIH3T3 cells expressing pHR-FUS-mCherry-Cry2, Cry2 existed intracellularly as a monomer in the absence of blue light stimulation. However, upon photo stimulation, Cry2 formed a dimer, clustered through LC domain-mediated multipoint interactions, and formed intracellular droplets. Further strong phototropic stimulation resulted in the formation of a phase-separated, gel-like structure. Once formed, the gel-like structures did not re-dissolve after cessation of light stimulation, but became irreversible aggregates. The monomer, dimer, and gelation spectra are shown in Fig. The spectral characteristics corresponding to each phase were as follows: (1) monomer: single peak (610 nm), (2) dimer: redshift of peak wavelength, from 610 to 612–620 nm, and (3) gelation: broadening of the spectrum and presence of bimodality (Figure 2b). Extraction of monomers, dimers, and gelled mCherry by spectral discrimination Time-lapse images were analyzed using the spectra shown in Figure 2b. These images were acquired over 12 cycles at 30-second intervals. Time 1 corresponded to the condition without light stimulation, while Times 2–12 involved blue light stimulation (0.03 mW/time). Without light stimulation (Time 1), mCherry fluorescence was diffusely present throughout the cytoplasm (red). After 60 s of light stimulation (Time 3, 0.06 mW), dots of dimers (green) were observed in the nucleus. The dimer region increased with increasing exposure. Further increase in exposure resulted in a gelatinized region after 240 s (Time 9, 0.24 mW). Thereafter, the gelatinized area increased (Figure 3a). Time 12 monomer, dimer, and gelation separation images. Exposure energy was 0.33 mW. Upper row: Image of one cell, white rectangular area, gelatinized area. The gelatinized area formed the core, surrounded by the dimer area, with monomers localized outside of it. The monomer portions were clustered around the dimer portion, with the gelatinized portion as the core from the cytoplasmic uniform distribution. Lower row: Enlarged view of the upper white line square area: Gel area (p1–p2) Center white line: Line profile measurement area. The line profile analysis of mCherry fluorescence intensity revealed that the gelation core corresponded to the gelation component (Cyan), surrounded by the dimer component (green), while the monomer component (red) was distributed at the periphery (Figure 3c). A schematic diagram of this organization is shown in Figure 3d. Discussion To experimentally determine whether an FP is likely to be a monomer or a dimer/oligomer, the most reliable method used is a combination of microscope-based optical and biochemical/physicochemical techniques. The following approaches are particularly effective: intensity-dependent analysis (comparing fluorescence intensity at different expression levels), fluorescence spectroscopy (Romani et al., 2010; Inomoto et al., 2007; Itoh & Osamra, 2007), fluorescence fluctuation spectroscopy / Number & Brightness analysis (Dunsin et al., 2018), fluorescence correlation spectroscopy / fluorescence cross-correlation spectroscopy (Yu et al., 2021), single molecule / stepwise photobleaching (Johan Hummert, et al. 2021), fluorescence Resonance Energy Transfer (FRET) / Fluorescence lifetime imaging microscopy-FRET (FLIM-FRET), fluorescence polarization (anisotropy) analysis, fluorescence recovery analysis, biochemical methods (chemical crosslinking + SDS-PAGE / Native PAGE), molecular weight measurement (SEC, SEC-MALS, and AUC), and mass spectrometry (crosslinking / native MS). We used a confocal laser microscopy wavelength analysis technique to reveal the differences between tag GFP and free GFP (Inomoto et al., 2007). The fluorescence spectrum (excitation or emission) of an FP is mainly affected by the environment surrounding its chromophores (Stepanenko et al., 2011). When monomers dimerize or oligomerize, (1) the distance and interactions between chromophores change, (2) minute structural changes in the protein lead to subtle shifts in pKa and electronic environment, and (3) FRET and self-quenching occur. As a result, the fluorescence wavelength may shift slightly, or the shape of the spectrum (shoulders and peak widths) may change. The change is often minimal, on the order of a few nanometers. The effects of other factors (pH, concentration, and ambient environment) are also likely to overlap. The spectral feature that directly indicates the differences between monomers and dimers is the wavelength shift, but this alone does not necessarily indicate dimerization (Möckel et al., 2019). Therefore, we analyzed this shift by combining it with morphological changes. The OptoDroplet method showed that optoIDR structures undergo photoactivated phase separation, but it is unclear whether the resulting clusters are liquid-like droplets or gel-like aggregates (Kato et al., 2012; Patel et al., 2015; Murakami et al., 2015; Weber & Brangwynne, 2012; Zhang et al., 2015; Lin et al., 2015; Molliex et al., 2015). In this report, we focused on the differences in the molecular morphology of the fluorescent probes, which allowed us to distinguish the properties of the resulting clusters using spectroscopic methods. In recent studies demonstrating that membrane-free cell organelles have partially solid-like properties (viscoelasticity), both nucleoli and stress granules have been reported to have a core-shell structure, with a gel-like core encased in a dynamic, liquid-like shell (Jain et al., 2016; Feric et al., 2016). Similarly, in the present study, the core of the gelatinized aggregates had a gel component, a dimer (droplet) component around it, and a monomer component further distributed outside of it. This method may provide a new way to elucidate the complexity of the intracellular aggregate formation process. Conclusion Microspectroscopic analysis successfully distinguished droplet formation and gel formation in the LLPS phenomenon of the optoDroplet method by recognizing differences in the three-dimensional structure of fluorescent molecules. Since abnormal protein aggregation like FUS is linked to neurodegenerative diseases, this method for studying LLPS mechanisms will support more detailed disease analysis and aid in developing new treatment technologies. This method is suggested to contribute to the elucidation of cell death mechanism caused by intracellular aggregate formation in neurodegenerative diseases. Authors’ Contributions J.I., N.F, S.K., M. I., N.S., and E.N. performed the experiments and analyzed the data. N.F generated NIH3T3 cells. J.I. and E.N designed the study and wrote the manuscript. All authors agree to be accountable for the content of the work. Figure legends Figure 1: Droplet formation process and filtration observation of wild type-mCherry-Cry2WT and FUS-mCHerry-Cry2WT by light stimulation a) Droplet formation after light stimulation of HN3T3 cells expressing pHR-mCherry-Cry2WT (wild type). Upper row: wild-type mCherry -Cry2WT spectrum with a single peak at 610 nm. Lower row: Time-lapse drawing with wavelength components in the wavelength range 602–620 nm. Time lapse: 12 cycles at 30 s intervals; time1; light off stimulation, time2-12; light stimulation (0.03mW/time). No droplet formation is observed in the wild type after light stimulation. b) Light-stimulated droplet formation in NIH3T3 cells expressing pHR-FUS-mCherry-Cry2WT. Upper row: Spectrum of pRH-FUS-mCherry-Cry2WT (a single peak at 610 nm). Lower row: Time-lapse drawing with wavelength components in the wavelength range 602–620 nm. Time lapse: 12 cycles at 30 s intervals; time1 light stimulation off, time2-12 light stimulation (0.03 mW/time). In NIH3T3 cells expressing pHR- FUS-mCHerry-Cry2WT with FUS placed in the LC domain, droplets appeared upon light stimulation (0.06 mW). Since the components are extracted by a bandpass filter, droplet shape (droplet) can be distinguished, but it is difficult to distinguish monomeric, dimeric, and trimeric components. Figure 2: Schematic diagram of LLPS a) Wild-type pHR-mCherry-Cry2WT remains in a state of weak bimolecular complex formation and does not form clusters, suggesting a reduced ability to interact with itself and a limited ability to self-associate, even after photo stimulation. Spectrum of wild-type pHR-mCherry-Cry2WT by spectroscopy with a single peak at 610 nm. b) pHR-FUS-mCherry-CRY2WT forms droplets upon light stimulation. This may be due to the interaction between FUS-LC domain molecules. When light stimulus is interrupted, the droplet resolves and returns to its original monomer. Upon further strong light stimulation, multiple intracellular aggregates (gelation) form along with the presence of droplets. Spectroscopy spectrum of pHR-FUS-mCherry-Cry2WT in monomer, dimer, trimer, or gelation states. Spectra displacement can be seen due to the difference in molecular structure. Figure 3: Extraction analysis of monomer, dimer, trimer, or gelation of FUN-mCherry-Cry2WT expressing NIH3T3 cells by spectroscopy Time-lapse separation images of monomer, dimer, trimer, or gelation using separation spectrum in Figure 2b. Time 1; light stimulation off, time 2–12; light stimulation on 0.03 mW/time). Red color: monomer, green color: dimer or trimer, cyan color: gelation. a) Time 1–2: monomer (red) only, no droplet formation, droplet appearance (green) at Time 3: number of droplets and droplet size increase with time, Time 6: gelatinization appears in part, Time 7 and after: gelatinization is accelerated. b) Upper row: Images of monomer, dimer, or trimer, and gelation separation at Time 12. Lower row: Enlarged image of the upper square. Square gelation area (p1 to p2). c) Line profile of p1 to p2 area. d) Schematic diagram of part C. 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JBC 294. 14823-14835. doi: 10.1074/jbc.REV119.007895 Supplementary Material File (figure 1.tif) Download 2.02 MB File (figure 2.tif) Download 1.16 MB File (figure 3.tif) Download 1.78 MB Information & Authors Information Version history V1 Version 1 27 October 2025 Copyright This work is licensed under a Non Exclusive No Reuse License. Keywords confocal laser microscopic spectroscopy liquid-liquid phase separation (llps) optodroplet Authors Affiliations Johbu Itoh Tokai Daigaku - Isehara Campus View all articles by this author Natsuko Fujii Tokai Daigaku - Isehara Campus View all articles by this author Saori Kohara Tokai Daigaku - Isehara Campus View all articles by this author Masatoshi Ito Sei Marianna Ika Daigaku View all articles by this author Naoto Suzuki Tokai Daigaku - Isehara Campus View all articles by this author Eiichiro Nagata [email protected] Tokai Daigaku - Isehara Campus View all articles by this author Metrics & Citations Metrics Article Usage 276 views 161 downloads .FvxKWukQNSOunydq8rnd { width: 100px; } Citations Download citation Johbu Itoh, Natsuko Fujii, Saori Kohara, et al. An Innovative Method for Evaluating Optical Properties of Liquid-Liquid Phase Separations using Laser Scanning Microscopic Spectroscopy. Authorea . 27 October 2025. 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