Self‐assembly of multi‐component mitochondrial nucleoids via phase separation

Article23 February 2021free access Source DataTransparent process Self-assembly of multi-component mitochondrial nucleoids via phase separation Marina Feric Marina Feric orcid.org/0000-0001-7531-9305 National Cancer Institute, NIH, Bethesda, MD, USA National Institute of General Medical Sciences, NIH, Bethesda, MD, USA Search for more papers by this author Tyler G Demarest Tyler G Demarest National Institute on Aging, NIH, Baltimore, MD, USA Search for more papers by this author Jane Tian Jane Tian National Institute on Aging, NIH, Baltimore, MD, USA Search for more papers by this author Deborah L Croteau Deborah L Croteau National Institute on Aging, NIH, Baltimore, MD, USA Search for more papers by this author Vilhelm A Bohr Vilhelm A Bohr orcid.org/0000-0003-4823-6429 National Institute on Aging, NIH, Baltimore, MD, USA Search for more papers by this author Tom Misteli Corresponding Author Tom Misteli [email protected] orcid.org/0000-0003-3530-3020 National Cancer Institute, NIH, Bethesda, MD, USA Search for more papers by this author Marina Feric Marina Feric orcid.org/0000-0001-7531-9305 National Cancer Institute, NIH, Bethesda, MD, USA National Institute of General Medical Sciences, NIH, Bethesda, MD, USA Search for more papers by this author Tyler G Demarest Tyler G Demarest National Institute on Aging, NIH, Baltimore, MD, USA Search for more papers by this author Jane Tian Jane Tian National Institute on Aging, NIH, Baltimore, MD, USA Search for more papers by this author Deborah L Croteau Deborah L Croteau National Institute on Aging, NIH, Baltimore, MD, USA Search for more papers by this author Vilhelm A Bohr Vilhelm A Bohr orcid.org/0000-0003-4823-6429 National Institute on Aging, NIH, Baltimore, MD, USA Search for more papers by this author Tom Misteli Corresponding Author Tom Misteli [email protected] orcid.org/0000-0003-3530-3020 National Cancer Institute, NIH, Bethesda, MD, USA Search for more papers by this author Author Information Marina Feric1,2, Tyler G Demarest3, Jane Tian3, Deborah L Croteau3, Vilhelm A Bohr3 and Tom Misteli *,1 1National Cancer Institute, NIH, Bethesda, MD, USA 2National Institute of General Medical Sciences, NIH, Bethesda, MD, USA 3National Institute on Aging, NIH, Baltimore, MD, USA *Corresponding author. Tel: +1 240 760 6669; E-mail: [email protected] The EMBO Journal (2021)40:e107165https://doi.org/10.15252/embj.2020107165 PDFDownload PDF of article text and main figures. Peer ReviewDownload a summary of the editorial decision process including editorial decision letters, reviewer comments and author responses to feedback. ToolsAdd to favoritesDownload CitationsTrack CitationsPermissions ShareFacebookTwitterLinked InMendeleyWechatReddit Figures & Info Abstract Mitochondria contain an autonomous and spatially segregated genome. The organizational unit of their genome is the nucleoid, which consists of mitochondrial DNA (mtDNA) and associated architectural proteins. Here, we show that phase separation is the primary physical mechanism for assembly and size control of the mitochondrial nucleoid (mt-nucleoid). The major mtDNA-binding protein TFAM spontaneously phase separates in vitro via weak, multivalent interactions into droplets with slow internal dynamics. TFAM and mtDNA form heterogenous, viscoelastic structures in vitro, which recapitulate the dynamics and behavior of mt-nucleoids in vivo. Mt-nucleoids coalesce into larger droplets in response to various forms of cellular stress, as evidenced by the enlarged and transcriptionally active nucleoids in mitochondria from patients with the premature aging disorder Hutchinson-Gilford Progeria Syndrome (HGPS). Our results point to phase separation as an evolutionarily conserved mechanism of genome organization. SYNOPSIS A dedicated set of proteins packages mitochondrial DNA (mtDNA) into nucleoprotein complexes to form mitochondrial nucleoids (mt-nucleoids). Here, mt-nucleoids are found to behave as biomolecular condensates, assembled via interactions between mtDNA and the main packaging protein TFAM. The mitochondrial genome is organized by phase separation. TFAM and mtDNA combine to form viscoelastic, multiphase droplets in vitro. TFAM surfactant behavior is mediated by its N-terminus favoring binding with DNA and its C-terminus preferentially interacting with mt-nucleoid associated proteins. Mitochondrial nucleoids exhibit similar phase separation behavior in vivo. Coalescence and enlargement of mt-nucleoids occur upon loss of mitochondrial homeostasis and in prematurely aged cells and may be associated with mitochondrial dysfunction. Introduction Mitochondria are the major sites of cellular energy production through oxidative phosphorylation and generation of ATP. Within each mammalian cell, mitochondria contain several hundred copies of a 16.6 kb double-stranded circular genome (Chen & Butow, 2005; Gustafsson, Falkenberg et al, 2016). Although small in size, the mitochondrial genome is gene dense, encoding essential polypeptides involved in mitochondrial respiration and oxidative phosphorylation (Burger, Gray et al, 2003). Unlike the nuclear genome, mtDNA is not organized by histones, but is packaged by a distinct set of proteins into nucleoprotein complexes to form mt-nucleoids (Chen & Butow, 2005; Kukat & Larsson, 2013). These structures are typically uniformly ~100 nm in size, each containing 1–2 molecules of mtDNA, and are spatially separated throughout the mitochondrial network (Brown, Tkachuk et al, 2011; Kukat, Wurm et al, 2011). mt-nucleoids lack any delimiting membranes, yet act as discrete functional units involved in replication and transcription of the mitochondrial genome (Kukat & Larsson, 2013). The major mt-nucleoid packaging protein in human cells is the mitochondrial transcription factor A (TFAM) (Chen & Butow, 2005), which binds and compacts mtDNA in vitro into nucleoid-like structures under dilute conditions (Brewer, Friddle et al, 2003; Kaufman, Durisic et al, 2007; Farge, Mehmedovic et al, 2014; Kukat, Davies et al, 2015). TFAM contains two high mobility group (HMG) domains that each intercalate into the DNA double helix, bending the DNA strand to form a tight U-turn structure at promoter sequences (Ngo, Kaiser et al, 2011; Rubio-Cosials, Sydow et al, 2011). Moreover, TFAM can also form loops and can bind cross-strands of DNA without sequence specificity (Kukat et al, 2015), and these conformations are further stabilized by cooperative TFAM-TFAM interactions (Farge, Laurens et al, 2012). In human cells, TFAM is present at high enough concentrations to coat the entirety of the circular mtDNA, driving the compaction of mtDNA from ~5 µm in contour length to the ~100 nm mt-nucleoid (Gustafsson et al, 2016). Beyond the direct binding of TFAM to mtDNA, it remains largely unknown how the higher-order morphological features of the mitochondrial genome emerge, how they affect function, and how anomalies in the structure of mt-nucleoids may contribute to disease (Friedman & Nunnari, 2014). Maintenance of mt-nucleoid structure is linked to mitochondrial organization and function. Disruption of mitochondrial fusion and fission processes affects mt-nucleoid size, as seen in knockdown of mitochondrial fission GTPase, Drp1, which leads to clustering of mt-nucleoids into large assemblages in hyperfused mitochondria (Ban-Ishihara, Ishihara et al, 2013; Ishihara, Ban-Ishihara et al, 2015). Similarly, downregulation of the inner mitochondrial membrane protein Mic60/Mitofilin leads to disassembly of the mitochondrial contact site and cristae organizing system (MICOS), ultimately resulting in enlarged, spherical mt-nucleoids (Li, Ruan et al, 2016). Enlargement and remodeling of mt-nucleoids are also associated with cellular stress responses. For example, prolonged exposure to DNA intercalating agents leads to altered mt-nucleoid size distributions (Ashley & Poulton, 2009; Alán, Špaček et al, 2016), and viral infection results in aberrant sizes of mt-nucleoids (West, Khoury-Hanold et al, 2015). Given the oxidative environment within mitochondria (Balaban, Nemoto et al, 2005; Sun, Youle et al, 2016) and the absence of protective histone proteins, the mt-nucleoid-associated proteins have been hypothesized to contribute to mitochondrial genome integrity (Yakes & Van Houten, 1997; Cadenas & Davies, 2000). Mutations in mtDNA have direct physiological relevance as elevated mutation levels of mtDNA are associated with premature aging phenotypes (Trifunovic, Wredenberg et al, 2004; Kujoth, Hiona et al, 2005). Moreover, mtDNA mutations tend to accumulate over the course of normal aging (Bratic & Larsson, 2013; Sun et al, 2016) and even single point mutations in mtDNA can elicit a myriad of other disease phenotypes (Taylor & Turnbull, 2005). An emerging organizational principle of non-membrane bound cellular structures is phase separation (Hyman, Weber et al, 2014). Numerous ribonucleoprotein and nucleoprotein complexes spontaneously self-assemble into non-membrane bound cellular bodies, or biomolecular condensates, via phase separation (Banani, Lee et al, 2017). The canonical examples of RNA-protein bodies include the nucleolus in the nucleus (Brangwynne, Mitchison et al, 2011; Feric, Vaidya et al, 2016) as well as P-granules (Brangwynne, Eckmann et al, 2009) and stress granules in the cytoplasm (Molliex, Temirov et al, 2015; Guillén-Boixet, Kopach et al, 2020; Sanders, Kedersha et al, 2020; Yang, Mathieu et al, 2020). In addition, DNA-protein complexes can phase separate in the nucleus, such as the heterochromatin protein HP1α in the context of heterochromatin (Larson, Elnatan et al, 2017; Strom, Emelyanov et al, 2017), histones to form chromatin domains (Gibson, Doolittle et al, 2019; Sanulli, Trnka et al, 2019), or super-enhancers which form active transcriptional hubs (Sabari, Dall'Agnese et al, 2018). Here, we have explored the higher-order organizational principles of the mitochondrial genome. We demonstrate, based on in vitro and in vivo observations, that mt-nucleoids self-assemble via phase separation. We find that the major mt-nucleoid protein TFAM exerts its architectural role by promoting phase separation via weak, multivalent self-interactions to generate the multiphasic mt-nucleoid structure. We also demonstrate that aberrant mt-nucleoid size is a consequence of phase behavior and is associated with mitochondrial dysfunction in the context of premature aging. Our observations suggest phase separation is an evolutionarily conserved mechanism in genome organization. Results Enlarged mitochondrial nucleoids in vivo During the course of in-depth analysis of cellular morphological changes in a model system of aging, we noticed the presence of aberrant mitochondria and enlarged mt-nucleoids in skin fibroblasts from patients with Hutchinson–Gilford Progeria Syndrome (HGPS) (Fig 1). HGPS is a rare, invariably fatal premature aging disorder characterized by multi-tissue symptoms, including of bone, muscle, skin, and cardiovascular failure. The disease is caused by a point mutation in LMNA resulting in the production of progerin, a dominant negative form of the major architectural protein lamin A (Gordon, Rothman et al, 2014). In line with mitochondrial abnormalities associated with HGPS (Rivera-Torres, Acín-Perez et al, 2013; Xiong, Choi et al, 2016), we find that ~70% of advanced HGPS patient cells had a sub-population of mitochondria that were swollen, spherical in shape and isolated from the surrounding mitochondrial network compared to the typical tubular, elongated mitochondrial networks in control cells (Fig 1A and B; Appendix Fig S1E–K). The extent and number of enlarged mitochondria correlated with disease progression (Fig 1C; Appendix Fig S1K), and several chaperones and proteases of the mitochondrial unfolded protein response (UPRmt) including HSPD1 (mtHSP60), mtHSP10, mtHSP70, ClpP, and LONP1 were enriched in enlarged mitochondria, indicating that the altered mitochondrial morphology is associated with mitochondrial stress (Fig 1J–L; Appendix Fig S1R–W). Exogenous expression of progerin was sufficient to induce in wild-type cells an increase in the number of aberrant mitochondria that scaled with progerin expression (Appendix Fig S1X–AF). Figure 1. Enlarged mitochondrial nucleoids are prominent in a premature aging disease and can also arise from liquid-like fusion events under stress A, B. Maximum intensity projections of SIM images of fixed normal (A) and HGPS (B) human skin fibroblasts, where the mitochondria are labeled in magenta with MitoTracker Red, mtDNA in green with anti-DNA, and the nucleus in blue with DAPI. Scale bar = 5 µm. C. Bar graph quantifying the number of damaged mitochondria per cell based on high-throughput imaging of two wild type and four HGPS primary skin fibroblast cell lines. Bars and error bars represent averages ± SEM for n = 3 independent experimental replicates (each experimental replicate had 15 technical replicates each containing 5 fields of view, approximately 2,000–5,000 total cells for each cell line analyzed), where P-value for the ANOVA test statistic was P < 0.001. For individual pairs, **P < 0.01, ***P < 0.001. D–G. Three-dimensional views of normal mitochondria (D) annotated by white box in (B) and swollen mitochondria (F) annotated by yellow box in (B) and showing TFAM localization in red with anti-TFAM; the length of the box = 4 µm. (E, G) Normalized intensity distributions of mt-nucleoids labeled with anti-DNA and with anti-TFAM corresponding to images from (D) and (F), respectively. H, I. Time-course experiment of live HGPS cells under phototoxic conditions. Mitochondria were labeled with MitoTracker Deep Red (H, magenta), and mtDNA was labeled with PicoGreen (H,I, green). Scale bar = 2 µm. Arrow heads indicate pairs of mt-nucleoids that undergo liquid-like fusion events. J, K. Single z-slices of SIM images of UPRmt in mitochondria from fixed normal (J) and HGPS (K) human skin fibroblasts, where the UPRmt marker is gray-scale with anti-HSPD1 (mtHSP60), and mtDNA is in green with anti-DNA. Scale bar = 1.5 µm. L. Immunofluorescence quantification of UPRmt markers (ClpP, LonPI, mtHSP10, mtHSP60, mtHSP70) in all six primary cell lines reported as normalized intensity values in undamaged and damaged mitochondria from high-throughput confocal images, where n = 3 independent experimental replicates (each experimental replicate, containing three technical replicates, had a total of 150–600 cells for each cell line and UPRmt marker), and error bars are standard error. Download figure Download PowerPoint Analysis by high-resolution Structured Illumination Microscopy (SIM) imaging revealed the presence of enlarged mt-nucleoids in morphologically aberrant mitochondria of HGPS cells (Fig 1F and G). In these mitochondria, mt-nucleoids clustered together into structures that were considerably brighter and larger than the typically uniform mt-nucleoids of ~100 nm found in normal mitochondria (Fig 1D and E). Several mt-nucleoid markers, including mtDNA and the major mtDNA-packaging protein, TFAM, were locally enriched in the atypical mitochondria (Fig 1F and G; Appendix Fig S1O and P), while total TFAM and mtDNA levels were not altered in HGPS cells (Appendix Fig S1L–N). The formation of enlarged mt-nucleoids is an indication of mitochondrial damage since they could also be induced in vivo when we exposed primary skin fibroblasts to phototoxic stress (Minamikawa, Sriratana et al, 1999) (see Materials and Methods). Within ~10 min, neighboring mt-nucleoids dynamically fused and relaxed into spherical, droplet-like structures (aspect ratio ≈ 1, time scale ≈ 20 ± 10 s, Appendix Fig S1Q) > 100 nm in size and up to a few microns in diameter analogous to those seen in damaged, swollen mitochondria of HGPS cells (Fig 1H and I and Movies EV1 and EV2). Similar, but even more pronounced, fusion events were observed in the presence of the intercalator EtBr (Movies EV3 and EV4). The homotypic fusion events between neighboring mt-nucleoids are consistent with the behavior of coalescing liquid droplets (Hyman et al, 2014; Banani et al, 2017). We conclude that loss of mitochondrial homeostasis results in the inability of mitochondria to maintain mt-nucleoid size and leads to the coalescence of multiple proximal mt-nucleoids to form larger, membrane-less droplets of nucleoprotein complexes in a process that closely resembles the phase separation of many other biomolecular condensates (Hyman et al, 2014; Banani et al, 2017). TFAM phase separates in vitro into viscoelastic droplets To explore if phase separation drives mt-nucleoid assembly, we examined the ability of the major mt-nucleoid packaging protein TFAM to undergo phase separation in vitro. In line with phase separation behavior, TFAM formed spherical droplets in low-salt conditions and at protein concentrations of ≥ 5 μM TFAM establishing a second proteinaceous phase, separate from the dilute, aqueous phase (Fig 2A; Appendix Fig S2D). Droplet formation was reversible upon increasing salt concentration (Appendix Fig S2F) and was reduced upon 1,6 hexanediol treatment (Appendix Fig S2G). At 30–60 min post-mixing, droplets coarsened to sizes of ~1–5 μm and sedimented toward the bottom of the imaging chamber (Fig 2A). The TFAM concentrations required for phase separation in vitro were well within the estimated physiological range inside the mitochondria on the order of ~10 µM (see Materials and Methods) (Kukat et al, 2011). Figure 2. TFAM phase separates into droplets with slow dynamics, driven by many weak, multivalent interactions Phase diagram (left panel) of TFAM under various protein and salt concentrations, where gray dots indicate single/soluble phase, red dots signify two phases/droplets present. DIC image (middle) and maximum intensity projection (right) of TFAM-DyLight-594 droplets at 25 µM and 150 mM NaCl in 20 mM Tris–HCl, pH 7.5 30 min after mixing. Scale bar = 5 µm. FRAP using 488 and 561 nm light performed on a ~1 µm spot on TFAM droplet 30 min after mixing. Inset shows representative fluorescent image of TFAM-DyLight-594 pre-bleach, immediately post-bleach, and 25 min post-bleach. Scale bar = 2 µm. Values represent averages ± SD from n = 15 droplets. Aspect ratio of droplet shape as a function of time after contact for a representative droplet. Inset shows fusion images corresponding to the trace at t = 0, 2, 4, and 40 min. Scale bar = 2 µm. The partition coefficient of dextran-FITC into TFAM droplets as a function of dextran average hydrodynamic radius estimated from the molecular weight. Solid line is an exponential fit to the data, where the partition coefficient is y = 6 e - R h / 0.6 + 0.4 giving rise to l ~ 1 nm. Inset shows representative images showing localization of dextran-FITC for Rh ≈ 1, 2 and 25 nm. Scale bar = 10 µm. Values represent averages ± SD from n = 3 experiments (>20 droplets analyzed per condition for each experiment). Schematic diagram of mutants with HMG domains in red and intrinsically disordered regions in gray. Yellow and green lines indicate point mutations in L6 and no-dimer mutants, respectively. Phase diagram of mutants at 150 mM NaCl and 20 mM Tris–HCl, pH 7.5 for a range of protein concentrations. Fluorescent maximum intensity projections of mutants at 50 µM protein and 150 mM NaCl, 20 mM Tris–HCl, pH 7.5 within 30–60 min after mixing. Scale bar = 2 µm. Download figure Download PowerPoint We performed a series of biophysical assays on TFAM droplets to assess their material properties within the first 30–90 min after mixing. Photobleaching of a R ≈ 0.5 μ m spot revealed slow dynamics with a characteristic time scale of τ ≈ 6.5 ± 0.5 min , which corresponded to a diffusivity of ∼ 6 × 10 - 4 μ m 2 / S . Furthermore, recovery was incomplete, with an immobile fraction of 0.5 ± 0.2, indicative of viscoelastic behavior (Fig 2B, Movie EV5). Similarly, time-lapse images of TFAM droplets undergoing coalescence events also displayed correspondingly slow dynamics with time scales of τ = 4 ± 0.5 min, giving rise to an inverse capillary velocity of 80 ± 20 s/µm (Fig 2C). Although the droplets had the propensity to relax upon contact, the average aspect ratio upon fusion was 1.36 ± 0.04, which deviated from that of a sphere (AR = 1.0) (Appendix Fig S2H). Introduction of dextran-FITC of varying sizes as an inert probe to sample the physicochemical environment of the droplets demonstrated that small particles of ≤ 1 nm preferentially accumulated within the droplets, most likely due to electrostatic interactions between negatively charged FITC and positively charged TFAM (Appendix Fig S2C), while increasing probe size resulted in reduced partitioning (Fig 2D). These properties are indicative of a characteristic pore or mesh size of ~1 nm, suggesting the presence of an effective polymer meshwork forming among individual TFAM molecules with markedly slow internal arrangements and signatures of viscoelasticity. TFAM phase separation is driven by multivalent interactions TFAM contains two DNA-binding high mobility group (HMG) domains separated by a disordered linker domain and flanked by an intrinsically disordered C-tail, together forming a relatively flexible protein chain (Appendix Fig S2A) (Ngo et al, 2011; Rubio-Cosials et al, 2011). Additionally, TFAM is one of the most highly charged proteins in the mt-nucleoid (Appendix Fig S2C) and is primarily enriched in positive amino acids distributed throughout the length of the protein (Appendix Fig S2B). To dissect the molecular features of TFAM responsible for phase separation, a set of TFAM mutants was analyzed for phase behavior in vitro (Fig 2E). In phase separation assays in vitro, the HMGA domain alone failed to form the typical micron-sized droplets as seen with full-length TFAM, but assembled into small puncta near the diffraction limit even at high concentrations (Fig 2F and G; Appendix Fig S2I–L). Maintaining half of the protein, either by adding a linker to HMGA (HMGA + linker) or with the structurally analogous HMGB + C-tail mutant, restored droplet formation, albeit at higher saturation concentrations than full-length TFAM, suggesting that multivalency seen in the full-length protein lowers the barrier for phase separation and that the addition of a disordered domain to HMGA promotes phase separation (Fig 2F and G; Appendix Fig S2I–L). Loss of the disordered C-tail (ΔC-tail) did not affect the saturation concentration, but influenced the wetting behavior of the droplets on the glass coverslip as indicated by a decreased smoothness along the droplet perimeter (Fig 2F and G; Appendix Fig S2I and J). The change in wetting behavior indicates differences in interfacial tension arising from molecular interactions between the droplet phase and the surroundings. Removal of the HMGA domain (ΔHMGA), which leaves a single HMG domain flanked by two disordered sequences, also resulted in droplet formation, but at slightly higher saturation concentrations. Finally, introduction of six non-polar residues in the linker region (L6 mutant) (Ngo, Lovely et al, 2014) affected the bulk properties of the droplets as evidenced by the highly non-spherical morphologies, whereas inclusion of non-polar residues in the HMGA domain to prevent dimerization (no-dimer mutant) (Ngo et al, 2014) increased saturation concentrations and produced smaller droplets, underscoring the contribution of multivalent interactions in phase separation of TFAM (Fig 2F and G; Appendix Fig S2I–L). Taken together, these observations suggest a mechanism by which many weak interactions along a flexible backbone of TFAM allow for robust phase separation and that the disordered linker and C-tail provide flexibility of the biopolymer chain to promote phase separation into prominent droplets. Formation of TFAM-mtDNA multiphase condensates in vitro To examine the interplay of mtDNA and TFAM in phase-separated mt-nucleoids, we investigated the in vitro phase behavior of TFAM in the presence of mtDNA (Fig 3; Appendix Fig S3A and B). As expected, mtDNA (0–10 nM or equivalently 0–100 ng/µl) on its own did not phase separate, but only when combined with TFAM at concentrations that support phase separation (≥ 5 µM), mtDNA readily partitioned into droplets. Importantly, the presence of mtDNA significantly affected droplet formation and morphology (Fig 3A; Appendix Fig S3C). Under initial mixing molar ratios of mtDNA/TFAM on the order of 1 × 10−5 – 1 × 10−3, which roughly correspond to the order of magnitude estimates of their physiological ratio (Kukat et al, 2011) (see Materials and Methods), TFAM and mtDNA readily associated into droplets together (Fig 3A; Appendix Fig S3C). At higher mtDNA/TFAM molar ratios (> 0.001), large, phase-dense droplets were no longer detected, potentially due to saturation behavior (Fig 3A; Appendix Fig S3C). However, at these high mtDNA/protein ratios, microscopic droplet formation could be restored upon addition of the crowder PEG (Appendix Fig S3D). For molar ratios of mtDNA/TFAM ≤ 6 × 10−4, the aspect ratio of the droplets notably increased with increasing ratio of mtDNA/TFAM molar concentrations (Fig 3B). For mtDNA/TFAM molar ratios < 3 × 10−4, the number of droplets (measured ~1 h after mixing) increased with increasing mtDNA/TFAM levels (Fig 3B inset), suggesting that mtDNA can potentiate droplet formation under those conditions, possibly acting as a nucleating agent and paralleling how RNA drives phase separation when added to RNA-binding proteins (Lin, Protter et al, 2015). ssDNA, dsDNA, and RNA as well as free nucleotides (dNTPs) also partitioned and supported TFAM droplet formation (Appendix Fig S3E–T). Interestingly, ssDNA and dsDNA, considerably longer than the 16.6 kB mtDNA and without any sequence specificity to mtDNA, resulted in even more irregular droplet morphologies (Appendix Fig S3E–T). We conclude that the addition of long, polymerized strands of DNA, irrespective of sequence, leads to favorable interactions between TFAM and DNA, thereby affecting the emergent droplet behavior (Appendix Fig S3E–V). While the aspect ratio of droplet morphologies did indeed vary across conditions tested, the structures that formed were invariably phase-dense and > 1 µm in diameter, corroborating that the TFAM-mtDNA in vitro system consistently exhibited phase separation as opposed to other modes of DNA compaction, such as salt-induced condensation or polymer collapse. These findings demonstrate that the material properties of the droplets depend on DNA/TFAM composition, where increasing DNA leads to more irregular droplet morphologies. Figure 3. TFAM and mtDNA form heterogenous droplets whose relaxation timescale is set by mtDNA in vitro A. Phase diagram of mtDNA versus TFAM denoting single/soluble phase (gray) or two phases/droplets (green). Each point on the phase diagram representing a unique DNA and protein concentration was measured from n = 2–12 independent experiments. Black solid line delineates deduced phase boundary. B. Aspect ratio as a function of dimensionless concentration (molar concentration mtDNA/molar concentration TFAM). Values represent binned mtDNA/TFAM conditions as measured in (A), and error bars are SEM. Green dashed line is an exponential fit to the data where y = 1.5 – 0.2 e ( − x / ( 3.8 × 10 − 4 ) ) . Inset: number of TFAM-mtDNA droplets per field of view as a function of dimensionless concentration. Values represent binned conditions from mtDNA/TFAM conditions measured in (A) and error bars are SEM. Green dashed line is an exponential fit to the data where y = 5,600 – 2,100e(−x/2.1). C–G. SIM images of droplets 30 min after mixing with various amounts of mtDNA: 0 nM (C), 0.1 nM (D), 1 nM (E), 4 nM (F), and 10 nM (G). Top row is of TFAM-DyLight-594 (red), middle row is of mtDNA-Alexa 488 (green) and bottom row is the merged image. Scale bar = 2 µm. H, I. FRAP experiments on TFAM-mtDNA droplets at 25 µM TFAM (H, red) and 10 nM mtDNA (I, green). Scale bar = 1 µm. J. FRAP recovery curve showing intensity as a function of time for TFAM (red) and mtDNA (green). Black line are single exponential fits, where mobile fraction ≈ 0.60 and characteristic recovery time t ≈ 350 ± 30 s for TFAM (y = 0.73–0.38exp(-t/350)), and recovery times t >> 15 min for mtDNA (error = 95% CI). Values represent averages ± SD from n = 16 droplets. Download figure Download PowerPoint To probe how mtDNA localizes within the droplets, we performed SIM imaging on TFAM-mtDNA droplets containing increasingly higher concentrations of mtDNA. We find that mtDNA is not uniformly distributed, but de-mixes from TFAM within the droplet (Fig 3C–G), consistent with multiphase behavior seen in other multi-component phase separating systems, and is reminiscent of the dense fibrillar component and fibrillar center of the nucleolus (Feric et al, 2016). Similar multiphase organization was observed with both ssDNA and dsDNA, but not with dNTPs nor RNA (Appendix Fig S3J–S). To characterize the dynamics of multiphase TFAM-mtDNA droplets, fluorescence recovery after photobleaching (FRAP) demonstrated that TFAM was able to diffuse within TFAM-mtDNA droplets with similar recovery behavior as in pure TFAM droplets (Fig 3H and J; Movie EV6). In contrast, on these timescales, mtDNA within the droplets remained strikingly immobile (Fig 3I and J; Movie EV6), suggesting that the mtDNA molecules within the TFAM-mtDNA droplets determine the time scale for the bulk properties of the droplets, while also explaining the observed non-spherical shapes at high mtDNA/TFAM ratios. The FRAP data suggest that these droplets represent kinetically arrested states arising from the slow dynamics associated with mtDNA inside the droplet, and as such, the