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Biomolecular condensates create phospholipid-enriched microenvironments

Nature Chemical Biology volume 20, pages 302–313 (2024)Cite this article

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Abstract

Proteins and RNA can phase separate from the aqueous cellular environment to form subcellular compartments called condensates. This process results in a protein–RNA mixture that is chemically different from the surrounding aqueous phase. Here, we use mass spectrometry to characterize the metabolomes of condensates. To test this, we prepared mixtures of phase-separated proteins and extracts of cellular metabolites and identified metabolites enriched in the condensate phase. Among the most condensate-enriched metabolites were phospholipids, due primarily to the hydrophobicity of their fatty acyl moieties. We found that phospholipids can alter the number and size of phase-separated condensates and in some cases alter their morphology. Finally, we found that phospholipids partition into a diverse set of endogenous condensates as well as artificial condensates expressed in cells. Overall, these data show that many condensates are protein–RNA–lipid mixtures with chemical microenvironments that are ideally suited to facilitate phospholipid biology and signaling.

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Fig. 1: A method for measuring condensate metabolomes.
Fig. 2: Phospholipids are enriched in condensates.
Fig. 3: Phospholipid hydrophobicity is important for condensate partitioning.
Fig. 4: Phospholipids partition into diverse types of condensates.
Fig. 5: Phospholipids can alter the quantity, size and morphology of MED1 condensates.
Fig. 6: Phospholipids are enriched in multiple condensates in cells.

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Data availability

Metabolomics data are publicly available at the NIH Common Fund’s National Metabolomics Data Repository website, the Metabolomics Workbench (https://www.metabolomicsworkbench.org), where it has been assigned project ID PR001509. The data can be accessed directly at https://doi.org/10.21228/M8N71K. Due to the size and lack of available condensate imaging databases, raw imaging data are available upon request to the corresponding author. Source data are provided with this paper.

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Acknowledgements

We thank all members of the Jaffrey laboratory for comments and suggestions. We thank S. Mitschka for producing the illustrations in Fig. 1b and the graphical abstract image. We also thank the Bio-Imaging Resource Center of Rockefeller University for assistance in performing confocal imaging (RRID:SCR_017791). This work was supported by the National Institutes of Health grants R35NS111631 and R01CA186702 (S.R.J.); R01AR076029, R21ES032347 and R21NS118233 (Q.C.) and P01 HD067244 and support from the Starr Cancer Consortium I13-0037 (S.S.G.).

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Author notes

  1. These authors contributed equally: Dawson Miller, Nabeel Attarwala.

Authors and Affiliations

  1. Department of Pharmacology, Weill Cornell Medical College, Cornell University, New York, NY, USA

    Jason G. Dumelie, Qiuying Chen, Dawson Miller, Nabeel Attarwala, Steven S. Gross & Samie R. Jaffrey

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  1. Jason G. Dumelie
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Contributions

J.G.D. and S.R.J. conceived the project and designed the experiments, unless otherwise stated. All authors contributed to the design of the condensate metabolomics experiments, and Q.C. and S.S.G. designed the LC–MS experiment and metabolite quantification following metabolite recovery from the input, aqueous and condensate fractions. D.M. and N.A. performed the LC–MS experiment, and Q.C. and D.M. quantified LC–MS peaks. J.G.D. performed all other experiments and analyses. J.G.D. and S.R.J. wrote the manuscript. The manuscript was read and approved by all authors.

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Correspondence to Samie R. Jaffrey.

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S.R.J. is scientific advisor to, and owns equity in, 858 Therapeutics. The remaining authors declare no competing interests.

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Nature Chemical Biology thanks Michal Holcapek and the other, anonymous, reviewer(s) for their contribution to the peer review of this work.

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Extended data

Extended Data Fig. 1 Developing a novel method for measuring condensate metabolomes.

a, MED1 and HNRNPA1 condensate FRAP imaging. mCherry-tagged MED1 (30 μM) or HNRNPA1 condensates (30 μM) were formed in LC-MS-compatible buffer by RNA addition (150 nM). After 10 min (25 °C), condensate sub-regions (indicated by dashed circle) were photobleached. Confocal images at the indicated intervals before and after photobleaching are shown. Scale bar, 5 μm. b, Quantification of MED1 and HNRNPA1 FRAP. Fluorescence was monitored at 4 s intervals before and after photobleaching. Fluorescence recovery (y-axis) was calculated at each timepoint as the percent of fluorescence recovered relative to observed fluorescence before photobleaching. This value was normalized using an unbleached region in the condensate. Lines represent mean fluorescence recovery across MED1 (n = 8 condensates) and HNRNPA1 (n = 6 condensates). c, Nucleocapsid condensates (30 μM) were formed in LC-MS-compatible buffer. NaCl (5 M) was added to the sample (final concentration 500 mM), which resulted in condensate depletion in 1 min (bottom). Scale bar, 5 μm (n = 3). d, Measurement of protein concentration in each fraction using mCherry fluorescence. Condensates were formed in LC-MS-compatible buffer, centrifuged and fractions were collected. Protein levels were determined based on mCherry fluorescence in equal amounts of each fraction. Median condensate enrichment was 22-fold for nucleocapsid (p = 0.002388), 14-fold for MED1 (p = 0.0006842) and 30-fold for HNRNPA1 (p = 0.029). Individual measurements are plotted as dots, bars represent means. Error bars indicate s.e.m. *p < 0.05 two-sided paired t-test (n = 3 per protein). e, Measurement of RNA concentration in each fraction. RNA was quantified from fractions in d. Median condensate enrichment was 30-fold for nucleocapsid (p = 0.01273), 106-fold for MED1 (p = 0.0315) and 68-fold for HNRNPA1 (p = 4.219e-05). Dots represent individual measurements, bars represent means. Error bars indicate s.e.m. *p < 0.05, two-sided paired t-test (n = 3 per protein, except MED1 top fraction where n = 2).

Source data

Extended Data Fig. 2 Processing and assessing quality of condensate metabolomics data.

Left, To determine whether thresholds needed to be imposed to remove low-abundant metabolites, median log2-metabolite ion counts (x-axis) were compared with the variation in metabolite ion counts (standard deviation in log2 counts, y-axis) across input samples. Based on the high variability of metabolites with <1000 median counts, metabolites with <1000 median counts per input sample were removed. High variation in input samples makes it difficult to calculate accurate enrichment scores, therefore metabolites with standard deviation log2(ion counts)>2.5 were also removed (n = 9). Thresholds are indicated with solid lines. Middle, To assess whether the short heating disruption step (2 min, 65 °C) during metabolite extraction from condensate, aqueous and input fractions impacts the measured condensate metabolome, we compared MED1 condensate metabolomes in the presence or absence of that step. The median log2-fold enrichment of phospholipids (blue, n = 43), lysophospholipids (green, n = 11), fatty acids (black, n = 14) and all other metabolites (orange, n = 210) for MED1 condensate metabolomics experiments including the heating step (x-axis) is plotted against the enrichment in the absence of the heating step (y-axis). Measured metabolite condensate enrichment in experiments in the presence or absence of the heating step are correlated (r = 0.93, Pearson’s correlation coefficient). Thus this step had minimal impact on MED1 metabolite enrichment measurements (n = 6 for samples with heat step, n = 3 for samples without heat step, n = 278 metabolites). Right, Evaluating the timing of metabolite addition to condensates. In most experiments, metabolites were added to proteins prior to condensate formation. Here we added the metabolite extract after condensate formation and 2 min prior to centrifugation. The median log2-fold enrichment of phospholipids (blue, n = 43), lysophospholipids (green, n = 11), fatty acids (black, n = 14) and all other metabolites (orange, n = 210) is correlated between experiments where metabolite extract was added before (x-axis) or after (y-axis) condensate formation (r = 0.92, Pearson’s correlation coefficient, n = 6 samples with metabolite extract added before, n = 3 added after, n = 278 metabolites).

Source data

Extended Data Fig. 3 The role of RNA in metabolite condensate partitioning.

a, To assess whether changes in generic phage RNA concentration alter condensate metabolomes, we compared nucleocapsid condensate metabolomes after varying the added RNA concentration. Median log2-fold enrichment of phospholipids (blue, n = 51), lysophospholipids (green, n = 14), fatty acids (black, n = 21) and all other metabolites (orange, n = 257) in the nucleocapsid condensate fraction relative to input sample with 150 nM RNA (x-axis) was plotted against median log2-fold enrichment with no RNA (y-axis; left) or 600 nM RNA (y-axis; right). Enrichment with 150 nM RNA correlates with enrichment in no RNA (r = 0.78, Pearson’s correlation coefficient) and 600 nM RNA (r = 0.85, Pearson’s correlation coefficient) samples. No metabolites were significantly different between 0 and 150 nM or 150 and 600 nM RNA samples (FDR > 0.1 for all metabolites, two-sided paired t-test with Benjamini-Hochberg adjustment). N = 3 for 150 nM RNA sample, n = 2 for other samples. b, To determine whether RNA concentration alters fatty acyl-dependent phospholipid partitioning, we compared the median log2-fold condensate enrichment of phospholipids (n = 51), lysophospholipids (containing one fatty acyl moiety, n = 14) and glycerophosphoryl head groups (for example glycerophosphocholine, n = 3) in the condensate fraction of nucleocapsid with different RNA concentrations. Phospholipids and lysophospholipids are enriched relative to head groups in condensates formed with 0 nM (green; p = 0.004105, phospholipids; p = 0.002941, lysophospholipids), 150 nM (purple; p = 0.004105, phospholipids; p = 0.002941, lysophospholipids, also shown in Fig. 3a) or 600 nM (black; p = 0.004105, phospholipids; p = 0.002941, lysophospholipids) RNA. Individual metabolites are represented by dots, lines in violin plots represent quartiles. *p < 0.05 **p < 0.005, two-sided Wilcoxon rank-sum test (n = 3 for 150 nM RNA sample, n = 2 for other samples). c, A heat map was used to further analyze whether RNA concentration impacts phospholipid enrichment in nucleocapsid condensates. Each glycerophosphoryl head group, lysophospholipid and phospholipid’s median log2-fold enrichment (blue indicates condensate de-enrichment and red indicates enrichment) is plotted for nucleocapsid condensates formed with 0 (left), 150 nM (middle) or 600 nM (right) RNA. Glycerophosphoryl head groups are sorted alphabetically, while lysophospholipids and phospholipids are sorted by the sum of carbons in their fatty acyl moieties after being grouped by head-group as demarked (y-axis).

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Extended Data Fig. 4 The role of fatty acyl moieties in phospholipid partitioning.

a, Individual phospholipid enrichment in condensates. To examine whether phospholipid head groups and fatty acyl moieties impact enrichment in condensates, we used a heat map. Each glycerophosphoryl head group, lysophospholipid and phospholipid’s median log2-fold enrichment (blue indicates condensate de-enrichment and red indicates enrichment) is plotted for nucleocapsid (left), MED1 (middle) and HNRNPA1 (right) condensates. Glycerophosphoryl head groups are sorted alphabetically, while lysophospholipids and phospholipids are sorted by the sum of carbons in their fatty acyl moieties after being grouped by head-group as demarked on the y-axis. We observe that longer phosphatidylethanolamines are less enriched in condensates for all three proteins, with similar phenomena observed with other head groups for specific proteins. This suggests that fatty acyl moiety length may reduce enrichment at least with some head groups and in some condensates (n = 3 for 150 nM RNA sample, n = 2 for other samples). b, Phospholipid fatty acyl moiety length is inversely correlated with partitioning. To determine if fatty acyl moiety chain length also contributes to partitioning, the total number of carbons in the fatty acyl moieties of lysophospholipids and phospholipids (x-axis, n = 65) was plotted against the median log2-enrichment of that lipid (y-axis) in nucleocapsid (left), MED1 (center) or HNRNPA1 (right) condensates relative to input sample using our metabolomics data. The number of unsaturated bonds is indicated (right) for each lipid with red indicating no unsaturated bonds, blue indicating more than 5 unsaturated bonds, with a gradient representing intermediate numbers of unsaturated bonds. There is no apparent correlation between the level of saturation at a given fatty acyl moiety length and the level of enrichment. On the other hand, there is a noticeable decrease in enrichment of phospholipids with longer fatty acyl moieties (>36 carbons) for all three proteins (Spearman’s rho < −0.41, p < 0.005, two-sided Spearman’s test). N = 3 replicates per protein.

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Extended Data Fig. 5 The role of charge and hydrophobicity in phospholipid partitioning.

a, Chemical library used for studying the effect of length and headgroup on phospholipid enrichment in MED1 condensates. b, Condensate metabolome experiments were performed using the chemical library from d rather than liver metabolite extract, with each molecule added to a final concentration of 100 nM (blue), 1 μM (green) or 10 μM (orange). The role of phospholipid tail length was assessed by plotting the combined tail length (x-axis) of each phosphatidylcholine against their log2-fold enrichment in MED1 condensates (y-axis). Choline glycerophosphoryl head group (carbons = 0) and the library’s lysophosphatidylcholine (carbons = 16) were also included. Results from each replicate are plotted as separate dots and crossbars represents means across replicates. Enrichment is only apparent with fatty acyl moiety chain length > 15. Combined fatty acyl moieties lengths of 16 and 18 have reduced enrichment at the lowest concentration, while phospholipids with longer fatty acyl moieties (>23) have higher enrichment at lower concentrations. Error bars indicate s.e.m. (n = 3). c, Condensate enrichment was compared between phospholipids with dipalmitoyl fatty acyl moieties, but different head groups using the library described in a. The log2-fold enrichment in MED1 condensates (y-axis) is plotted for each phospholipid (x-axis) when metabolites were added 100 nM (blue), 1 μM (green) or 10 μM (orange). Results of individual replicates are plotted as dots (black), while bars indicate the mean. Error bars indicate s.e.m. (n = 3). d, Net neutral charge phospholipids preferentially partition into MED1 and HNRNPA1 condensates. Phospholipids from the metabolomics datasets analyzed in Fig. 2 were grouped based on head group (x-axis): sphingomyelins (SM, n = 4), phosphatidylcholines (PC, n = 4), phosphatidylethanolamines (PE, n = 27), phosphatidylinositols (PI, n = 5), and phosphatidylserines (PS, n = 11). For each group, median log2-fold enrichment was plotted for nucleocapsid (purple), MED1 (blue; p = 0.02637, SM/PS; p = 0.02584, PE/PI; p = 0.0001876, PE/PS) or HNRNPA1 (green; p = 0.01587, SM/PI; p = 0.001465, SM/PS, p = 0.01587, PC/PI; p = 0.001465, PC/PS; p = 0.00149, PE/PI; p = 6.79e-06, PE/PS) condensates. Violin plots represent the distribution of median enrichment for each head group and lines demarcate quartiles. *p < 0.05, **p < 0.005, two-sided Wilcoxon rank-sum test (n = 3 condensate metabolomic experiments per protein).

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Extended Data Fig. 6 Phospholipids co-localize with condensates under diverse conditions.

a, Oregon Green phospholipid is enriched in nucleocapsid condensates under metabolomics buffer conditions. Nucleocapsid (30 µM, red) was combined with 2 µM Oregon Green dye (top, green) or phosphatidylethanolamine (bottom, green) in LC-MS-compatible buffer (50 mM NH4HCO3 pH 7.5, 50 mM NaCl, 1 mM DTT) and then generic phage RNA (150 nM) was added to promote condensate formation. Condensates were imaged after a 10 min incubation. A representative image is displayed for each condition. Oregon Green phosphatidylethanolamine, but not dye, colocalizes with each condensate. Scale bar, 5 μm. b, Quantification of a. The median ratio of mean fluorescent signal inside condensates to the mean signal outside condensates for Oregon Green dye (Dye, orange) or phosphatidylethanolamine (Dye-phospholipid, blue) in z-stacks 1-3 μm above the slide surface (y-axis) is plotted for each condition (x-axis). Oregon Green phosphatidylethanolamine is enriched relative to dye (p = 0.005083). Error bars indicate s.e.m. *p < 0.05, two-sided Welch’s t-test (n = 3 imaging experiments). c, Oregon Green phospholipid is enriched in nucleocapsid condensates in the absence of mCherry. To determine whether mCherry might drive phospholipid partitioning into nucleocapsid condensates, we asked whether Oregon Green phospholipids partition into nucleocapsid condensates that lack mCherry. Oregon Green phosphatidylethanolamine or dye (2 µM, green) was added to solutions of nucleocapsid (3 µM, grey-scale) in the presence of phage RNA (15 nM) in buffer (50 mM Tris pH 7.5, 140 mM KCl, 12 mM NaCl, 0.8 mM MgCl2, 5% PEG-8000). Nucleocapsid condensates were imaged after 10 min using phase contrast microscopy, while imaging Oregon Green with fluorescence microscopy. Representative images are shown for each condition and two in-frame nucleocapsid condensates are expanded below. Oregon Green phosphatidylethanolamine, but not dye, colocalizes with the nucleocapsid condensate suggesting that the mCherry-tag is not required for phospholipid partitioning. Scale bar, 5 μm (n = 3 for phosphatidylethanolamine samples, n = 2 for dye samples).

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Extended Data Fig. 7 Phospholipase treatment inhibits Oregon Green phosphatidylethanolamine condensate partitioning.

a, Phospholipase cleavage sites. Arrows indicate location of phospholipase cleavage. R indicates fatty acyl chains. b, Removing fatty acyl moieties from Oregon Green phosphatidylethanolamine depletes condensate enrichment. Oregon Green phosphatidylethanolamine (green, 2 μM) was pretreated with each of the indicated phospholipases, or vehicle, prior to being combined with nucleocapsid (red, 3 μM) and RNA (15 nM) in buffer lacking divalent cations (50 mM Tris pH 7.5, 140 mM KCl, 12 mM NaCl, 5 mM EDTA, 5% PEG-8000). Samples were imaged by fluorescence microscopy after a 10 min incubation. Pretreatment with any of the three phospholipases leads to reduced Oregon Green signal enrichment in condensates, indicating that both fatty acyl moieties are needed for condensate enrichment. EDTA inhibits the activity of both phospholipase A2 and D. EDTA (5 mM) was added, as indicated, during the phospholipid pretreatment with these phospholipases as controls. EDTA addition during phospholipase treatment with phospholipases A2 or D restored phospholipid condensate enrichment. Representative image are shown for each condition, scale bar, 5 μm. c, Quantification of b. The median ratio of mean fluorescence signal inside nucleocapsid condensates to the mean signal outside nucleocapsid condensates for Oregon Green phosphatidylethanolamine (blue) across z-stacks 1–3 μm above the slide surface was plotted for each enzymatic treatment. EDTA restores phospholipid enrichment for PLA2 (p = 0.002764) and PLD (p = 0.04016). Error bars indicate s.e.m. *p < 0.05, two-sided Welch’s t-test (n = 2).

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Extended Data Fig. 8 Phospholipids can alter condensates.

a, Oregon Green phosphatidylethanolamine did not alter MED1 condensate size in Fig. 5a. Microscopy images in the experiment performed in Fig. 5a were segmented using ImageJ RAST (see Methods). Particles > 0.1 µm2 were considered condensates. Median area of these condensates across five images/replicate (y-axis, blue) is plotted for Oregon Green dye or Oregon Green phosphatidylethanolamine (x-axis). Error bars indicate s.e.m. NS, not significant, two-sided Welch’s t-test (n = 6 imaging experiments). b,c, Quantification of the number (b) and size (c) of MED1 condensates after phosphoinositide addition. The median number of condensates/image (y-axis) and size of condensates (y-axis; p = 0.02776, PIP2) were quantified for each replicate for each type of phosphoinositide imaged in Fig. 5d (x-axis). Error bars indicate s.e.m. NS, not significant *p < 0.05, two-sided Welch’s t-test (n = 4 imaging experiments). d, Quantification of the number of large MED1 condensates in Fig. 5d. Particles with area > 1 µm2 were considered large condensates. The median number of large condensates/image across five images (y-axis) was plotted for each imaged condition (x-axis). More large condensates formed after PIP2 (p = 0.00783) or PIP3 (p = 0.01359) addition than PI addition. Error bars indicate s.e.m. *p < 0.05, two-sided Welch’s t-test (n = 4 imaging experiments). e, To determine if interactions between PIP3 phosphate groups and divalent cations affect MED1 condensate morphology, condensate formation was performed without (top) or with (middle) EDTA (5 mM), or with both EDTA and extra MgCl2 (bottom, final concentration 5.8 mM) for 15 min (25 °C). A representative image is displayed for each condition, with the region around one condensate expanded (right). Scale bar, 5 μm (n = 4 imaging experiments). f, Quantification of MED1 condensate shape changes in e. Irregularity of condensates was quantified by subtracting the measured condensate circularity (which is between 0 and 1) for samples in e from one. Median condensate irregularity across five images/replicate (y-axis, blue) was plotted for each condition (x-axis). MED1 condensates were more circular with EDTA than without EDTA (p = 0.005526) or with both EDTA and extra MgCl2 (p = 0.01472). Error bars indicate s.e.m. *p < 0.05, two-sided Welch’s t-test (n = 4 imaging experiments).

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Extended Data Fig. 9 Neomycin depletes anti-PIP2 and anti-PIP3 immunostaining.

a, Neomycin was used to test the specificity of anti-PIP2 immunofluorescence. Neomycin binds to the head groups of phosphorylated phosphoinositides. Cells were co-immunostained for PIP2 (green) and SON. Either neomycin (2 mg/ml) or vehicle (PBS) was added to both the pre-antibody incubation with 2% FBS and during the primary antibody incubation. PIP2 images are displayed with increased brightness for cytoplasmic granules since there is less PIP2 in the cytoplasm than the nucleus. A representative image is displayed for each immunostain. Scale bar, 5 μm (n = 2). b, Metaplot of PIP2 signal proximal to condensates imaged in a. PIP2 signal was measured in the area surrounding each condensate’s center. The median protein (red) and PIP2 (green) signal intensity across all condensate images is plotted. Each metaplot represents one representative experiment. Both the vehicle and neomycin treatments for each experiment were performed at the same time using the same imaging and display conditions. Neomycin treatment depletes the PIP2 signal observed in condensates (n = 2). c, To determine if anti-PIP3 immunofluorescence in condensates is specific to PIP3, neomycin was used to deplete anti-PIP3 immunofluorescence. Cells were co-immunostained for PIP3 (green) and G3BP1 as described in Fig. 6c. In the presence of neomycin, there is still significant amount of background signal across the cell, but the enrichment of PIP3 in stress granules and P bodies is no longer observed, suggesting that signal is specific. A representative image is displayed for each immunostain. Scale bar, 5 μm (n = 2). d, Metaplot of PIP3 signal proximal to condensates imaged in c. PIP3 signal was measured in the area surrounding each condensate’s center. The median protein (red) and PIP3 (green) signal (intensity) across all condensate images from an experiment is plotted for immunostains with or without neomycin (as indicated), with the distance from the center of each condensate indicated on the x-axis and y-axis. Both the vehicle and neomycin treatments for each experiment were performed at the same time using the same imaging and display conditions. PIP3 signal in condensates is depleted by neomycin (n = 2).

Extended Data Fig. 10 Variable anti-PIP2 and anti-PIP3 antibody signal in synthetic condensates formed by LAF-1 RGG domains.

a, Metaplot of PIP2 and PIP3 signal proximal to condensates imaged in Fig. 6e. The location of condensates was identified in a semi-automated manner. Anti-PIP2 (above) or anti-PIP3 (below) immunofluorescence was then measured in the area surrounding each condensate’s center. The median GFP-LAF1 (red, left) and anti-PIP2 or anti-PIP3 (green, right) signal intensity across all examined synthetic condensates (from n = 3 biological replicates) is displayed with the distance from the center of each condensate indicated on the x-axis and y-axis. Median PIP2 and PIP3 immunofluorescence is higher in synthetic condensates formed by LAF-1 RGG domains than in adjacent regions. The number of condensates examined is indicated in each row. b, PIP2 and PIP3 are not enriched in synthetic condensates within select cells. A synthetic, condensate-forming protein containing GFP and two copies of the C. elegans LAF-1 protein’s RGG domain was over-expressed from a plasmid. Immunofluorescence was performed as described in Fig. 6a, using antibodies against either PIP2 (top) or PIP3 (bottom). Approximately two-third of the cells containing LAF-1 condensates had condensates that did not co-localize with either phosphoinositide and these images are representative of those cells. A region containing condensates is highlighted by a white square and expanded in the adjacent image (right). Scale bar, 5 μm (n = 3 biological replicates).

Supplementary information

Supplementary Information

Supplementary Figs. 1–13, Tables 1–3 and source data for Supplementary Fig. 1.

Reporting Summary

Supplementary Data Set 1

Processed metabolomics data.

Supplementary Video 1

Nucleocapsid condensates move rapidly under buffer conditions used in condensate metabolomics assays. Fluorescence microscopy time-lapse images were performed to determine if nucleocapsid condensates were amenable to FRAP microscopy. Nucleocapsid (30 μM; red) condensates were formed in condensate metabolomics buffer (50 mM NH4HCO3 (pH 7.5), 50 mM NaCl and 1 mM DTT) by adding mouse liver metabolites (150 g liter–1 protein equivalent) and then generic phage RNA (150 nM). Imaging occurred after a 10-min incubation (25 °C), with images taken every 4 s using an inverted laser-scanning confocal microscope with a ×63 lens and immersion oil. The constant movement of nucleocapsid condensates prevented us from performing FRAP with these condensates (n = 2); scale bar, 5 μm.

Supplementary Video 2

More MED1 condensates settle on the bottom of wells with the addition of Oregon Green phospholipid than with the addition of Oregon Green dye. Condensates were formed containing MED1 (3 μM; red), generic phage RNA (15 nM) and either Oregon Green dye (left; 10 μM) or Oregon Green-tagged phosphatidylethanolamine (right; 10 μM). Condensate formation was stimulated by adding PEG-8000 (5%). Images from both samples were taken nearly simultaneously in adjacent wells under identical conditions, at the noted time (bottom left) after PEG-8000 addition, with a widefield fluorescence microscope using a ×60 lens with immersion oil. Images were taken just above the glass surface because once MED1 condensates settle on the glass surface of the well, they become immobile, making them easier to image over time. Images were deconvoluted as described in the Methods. Throughout the time-lapse images, more MED1 condensates settled on the glass surface with the addition of Oregon Green-tagged phosphatidylethanolamine than with the addition of Oregon Green-tagged dye. A representative time-lapse image is displayed (n = 3); scale bar, 5 μm.

Supplementary Video 3

PIP3 addition induces the formation of MED1 condensates with a different morphology and size than MED1 condensates formed with PI. Condensates were formed containing MED1 (3 μM; red), generic phage RNA (15 nM) and either PI (left; 10 μM) or PIP3 (right; 10 μM). Condensate formation was stimulated by adding PEG-8000 (5%). Images from both samples were taken nearly simultaneously in adjacent wells under identical conditions, at the noted time (bottom left) after PEG-8000 addition, with a widefield fluorescence microscope using a ×60 lens with immersion oil. Images were taken just above the glass surface because once MED1 condensates settle on the glass surface of the well, they become immobile, making them easier to image over time. Images were deconvoluted as described in the Methods. MED1 condensates are larger and less round after PIP3 addition than after PI addition throughout the time course. A representative time-lapse image is displayed (n = 3); scale bar, 5 μm.

Supplementary Video 4

Mg2+ is required for the unusual MED1 condensate shape induced by PIP3 addition. Condensates were formed containing MED1 (3 μM; red), generic phage RNA (15 nM) and PIP3. Condensate formation was stimulated by adding PEG-8000 (5%). Images from both samples were taken nearly simultaneously in adjacent wells using identical imaging conditions with a widefield fluorescence microscope using a ×60 lens with immersion oil. After 15 min, water (left) or 5 mM EDTA (right) was added. The time relative to EDTA addition is indicated in the bottom left. Images were deconvoluted as described in the Methods. MED1 condensates become more round after EDTA addition and are unaffected by the addition of a similar volume of water. This suggests that Mg2+ is necessary for the effect of PIP3 on MED1 condensates and that, despite their morphology, the PIP3-induced condensates can dynamically rearrange their morphology after removal of available divalent cations. Representative regions are shown (n = 3); scale bar, 5 μm.

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Dumelie, J.G., Chen, Q., Miller, D. et al. Biomolecular condensates create phospholipid-enriched microenvironments. Nat Chem Biol 20, 302–313 (2024). https://doi.org/10.1038/s41589-023-01474-4

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