Nanotubular Highways for Intercellular Organelle Transport

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Nanotubular Highways for Intercellular Organelle Transport Amin Rustom,1 Rainer Saffrich,2 Ivanka Markovic,3 Paul Walther,4 Hans-Hermann Gerdes1,5* Cell-to-cell communication is a crucial prerequisite for the development and maintenance of multicellular organisms. To date, diverse mechanisms of intercellular exchange of information have been documented, including chemical synapses, gap junctions, and plasmodesmata. Here, we describe highly sensitive nanotubular structures formed de novo between cells that create complex networks. These structures facilitate the selective transfer of membrane vesicles and organelles but seem to impede the flow of small molecules. Accordingly, we propose a novel biological principle of cell-to-cell interaction based on membrane continuity and intercellular transfer of organelles. Performing three-dimensional (3D) live-cell microscopy (1) in the presence of fluorescently labeled lectin wheat germ agglutinin (1), we observed ultrafine intercellular structures of cultured rat pheochromocytoma PC12 cells. These structures, referred to here as tunneling nanotubes (TNTs), had a diameter of 50 to 200 nm and a length of up to several cell diameters (Fig. 1, A to G). TNTs rarely displayed a branched appearance (Fig. 1C, arrow). Furthermore, they were stretched between interconnected cells attached at their nearest distance and did not contact the substrate (Fig. 1D). TNTs were also observed in human embryonic kidney (HEK) or normal rat kidney (NRK) cells (fig. S1). TNTs displayed a pronounced sensitivity to prolonged light excitation, leading to visible vibrations and rupture (fig. S2). Mechanical stress and chemical fixation also resulted in the rupture of many TNTs. However, trypsin-EDTA treatment did not disrupt TNTs (fig. S3). TNTs contained F-actin but not microtubules (Fig. 1E). Similar findings have been reported for cellular extensions termed cytonemes, first observed in the Drosophila wing imaginal disc (2). When we performed scanning electron microscopic (SEM) analysis, the stretched shape and structure of TNTs could be preserved, and their surface showed a seamless transition to the surface of both connected cells (Fig. 1F). Transmission electron microscopic (TEM) analysis changed the stretched morphology of TNTs into a bent Interdisciplinary Center of Neuroscience (IZN), Institute of Neurobiology, University of Heidelberg, INF 364, Heidelberg 69120, Germany. 2Otto-MeyerhoffZentrum, University of Heidelberg, INF 350, Heidelberg 69120, Germany. 3Institute of Biochemistry, Faculty of Medicine, University of Belgrade, Pasterova 2, Belgrade 11000, Yugoslavia. 4Electron Microscopy Facility, University of Ulm, Albert-Einstein-Allee 11, 89069 Ulm, Germany. 5Institute for Biochemistry and Molecular Biology, University of Bergen, Jonas Lies vei 91, Bergen 5009, Norway.

configuration presumably because of mechanical stress during sample preparation. However, serial sectioning showed that, at any given point along TNTs, their membrane appeared to be continuous with the membranes of connected cells (Fig. 1G). The peculiar morphology of TNTs raised the question of how they are generated. Over a 4-min period, a cell formed filopodia-like protrusions seemingly directed toward a

neighboring cell (Fig. 2, A and B). One protrusion made contact (Fig. 2C), which resulted in TNT formation (Fig. 2D, arrow) and the degeneration of remaining protrusions (Fig. 2, compare B and D). The number of TNTs increased during the first 2 hours after plating of single cells (fig. S4A). TNTs did not appear to be relics of incomplete cytokinesis (fig. S5). Thus, our data strongly suggest that TNTs are formed de novo. TNT formation was not an event restricted to pairs of cells but could lead to complex cellular networks (Fig. 2, E and F). Because TNTs were frequently found between diverging cells (Fig. 2F, black arrows), they also may exist between associated cells. After treatment with latrunculin-B, a substance that depolymerizes F-actin, no TNTs were detectable (fig. S4B), which suggested that actin-driven cellular protrusions participate in TNT formation. The existence of expanded intercellular networks prompted us to test whether TNTs can participate in cell-to-cell communication. Videomicroscopic analysis (1) revealed tubular or vesicular objects moving in one direction at a speed of 25.9 ⫾ 7.9 nm/s along given TNTs (Fig. 2, G and H). This phenomenon showed striking similarities to the demonstrated transfer of lipid containers between liposomes via phos-

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*To whom correspondence should be addressed. Email: [email protected]

Fig. 1. (A to D) Architecture of TNTs between cultured PC12 cells. Wheat germ agglutinin–stained PC12 cells were analyzed by 3D live-cell microscopy. Cells are connected via one (A) or several TNTs (B) with surrounding cells. Rarely, branched TNTs were observed [(C), arrow]. In (D) a selected (x-z) section obtained from a confocal 3D reconstruction is shown. (E) TNTs contain actin but no microtubules. Fixed PC12 cells were immunostained with an antibody against ␣-tubulin (green), phalloidine–fluorescein isothiocyanate (FITC) (red), and DAPI (blue). A single (x-y) section of a deconvolved 3D reconstruction is shown. The inset depicts the corresponding (x-z) section through the marked TNT (arrow). (F and G) Ultrastructure of TNTs. PC12 cells analyzed by SEM (F) or TEM (G) of consecutive 80-nm sections (G1, G2). For boxed areas, higher magnification images are shown (F1 to F3, G1, G2). Open arrowhead, secretory granule. Scale bars: (A to E), 15 ␮m; (F), 10 ␮m; (G), 2 ␮m; (F1 to F3, G1, G2), 200 nm.

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REPORTS Fig. 2. De novo formation of TNTs. Two hours after plating, PC12 cells were imaged by time-lapse bright-field microscopy. (A to D) Different stages of TNT formation. Selected frames of a video sequence (Movie S1, acquired at 2 frames per second over 4 min) are shown (A). One protrusion [(B), arrowhead] makes contact with a neighboring cell (C), which resulted in TNT formation [(D), arrow]. (E and F) TNTmediated network formation. Timelapse videomicroscopy (3D) shows that TNTs become visible between diverging cells (black arrows) and are formed de novo (white arrows). Shown are overlays of three selected z-sections each, displaying all detectable TNTs. Numbers refer to corresponding cells. (G and H) Unidirectional translocation of an object (arrowhead) along TNTs. Selected frames of a video sequence (Movie S2, acquired at 1 frame per min over 4 min) are shown. The arrow indicates the starting position in translocation. Time points of image acquisition are indicated. Scale bars, 20 ␮m.

Fig. 3. Transfer of soluble and membrane marker molecules between TNT-connected cells. (A and B) Transport of organelles. PC12 cells were stained with LysoTracker and analyzed by fluorescence videomicroscopy. Two selected frames of a video sequence (Movie S3, acquired at 1 frame per 2 s, over 4 min) are shown. Arrowheads mark LysoTracker-stained organelles; arrows indicate their starting position in translocation. Time points of image acquisition are indicated. (C to E) Partial colocalization of synaptophysin and myosin Va. Fixed PC12 cells analyzed by immunofluorescence microscopy with antibodies against synaptophysin [(C and E), green] or myosin Va [(D), green; (E), red] and tetramethyl rhodamine isothiocyanate (TRITC)–phalloidine [(C and D), red; (E), blue]. Open arrowheads indicate selected punctuated signals of both proteins, and the open arrow indicates their partial colocalization. Dashed lines indicate parts of the cell borders. (F to Q2) Transfer of EGFP fusion proteins. PC12 cells stained with CellTracker (blue) were mixed with cells transfected with synaptophysin-EGFP [syn-EGFP (F to I2)], EGFP-actin (J to M2), or farnesylated-EGFP [f-EGFP (N to Q2)] and cocultured for 24 to 48 hours. Cells were processed for immunocytochemistry with GFP-specific antibody (red) and phalloidine-FITC (green), and analyzed by 3D microscopy. Single (x-y) sections of deconvolved 3D reconstructions are shown as indicated. The numbers in the single-channel recordings (top rows) refer to the transfected cells (population 1) or CellTrackerstained cells (population 2). Boxed areas are magnified in (I1), (M1), (Q1), respectively. A second z-section for the same area is shown (I2, M2, and Q2). Arrows, TNTs; arrowheads, transferred marker proteins. Scale bars, 10 ␮m.

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pholipid bilayer nanotubes generated in vitro (3). After labeling acidic organelles with LysoTracker (1), we monitored fluorescent structures as they traveled unidirectionally inside TNTs (Fig. 3, A and B). Immunocytochemical analysis (1) revealed that synaptophysin, a marker for early endosomes and endosomederived, small synaptic-like microvesicles (SLMVs) (4), was present as discrete signals inside TNTs (Fig. 3C, open arrowheads). Myosin Va, a motor protein shown to facilitate organelle transport (5), was also present inside TNTs (Fig. 3D, open arrowheads) and partly colocalized with SLMVs (Fig. 3E, open arrow), which is consistent with an actin-dependent transport mechanism. To investigate whether membrane containers could be exchanged between TNT-connected cells, we analyzed the transport of synaptophysin fused to enhanced green fluorescent protein (EGFP) between two different cell populations. One population transfected with synaptophysinEGFP (population 1) was cocultured for 24 to 48 hours with a second population labeled with CellTracker (population 2). Organelles that stained positive for synaptophysin-EGFP could be detected selectively in those cells of population 2 that were connected via a TNT with synaptophysin-EGFP-expressing cells (Fig. 3, F to I2, arrowheads). To investigate whether cytoplasmic molecules could also be transferred between TNT-connected cells, we tested for the distribution of cytoplasmically expressed actin fused to EGFP. EGFP-actin could be detected in or near the actin cortex of TNT-connected cells of population 2 (Fig. 3, J to M2, arrowheads). These signals were similar to the patchy signals of EGFP-actin found in cells of population 1 (fig. S6). Patchy signals of EGFP-actin most likely represent prominent actin-rich foci referred to as “actin patches” (6, 7). The TNTdependent transfer of a soluble marker protein is consistent with the existence of membrane continuity, as suggested by the ultrastructural analyses. However, neither cytoplasmically ex-

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Fig. 4. TNT-based transfer of membrane components and dye-labeled organelles. (A to C) Intercellular membrane flow. PC12 cells stained with CellTracker were mixed with cells transfected with f-EGFP, cocultured for 48 hours, and analyzed by 3D videomicroscopy. Numbers in the single-channel recordings (B and C) refer to the transfected cells (population 1) or CellTrackerstained cells (population 2). The inset in (A) depicts the corresponding (x-z) section through the marked TNT (arrow). Arrowheads, staining derived from transferred f-EGFP. (D to E2) Induced transfer of f-EGFP. PC12 cells were transfected with f-EGFP and incubated for 12 hours. A selected f-EGFP–positive cell (labeled 1), connected via a TNT (arrow) with a nonexpressing cell (2), was microinjected with hyperosmotic solution and analyzed 10 min thereafter. Asterisks mark neighboring nonexpressing cells. For quantification, line profiles displaying gray value intensities [(E1 and E2), right] were performed as indicated (red arrows). Arrowheads, position of the cell border. (F and G) Unidirectional transfer of DiI-labeled organelles. PC12 cells were stained with DiI and analyzed by fluorescence videomicroscopy. A selected frame of three consecutive video sequences (Movies S4 to S6, acquired at 4 frames per s) showing DiI-labeled organelles within a TNT [(F), arrow] is shown. Broken lines indicate parts of the cell borders. Arrows 1 to 4 mark trajectories of four selected organelles (for details, see fig. S11). (H to I1) Intercellular organelle transfer correlates with the existence of a TNT connection. A mixed population of DiI- (red) and DiO-labeled PC12 cells (green) was plated and incubated for 12 hours. Pairs of TNT-connected cells identified by differential interference contrast microscopy [(H), arrow] were analyzed for organelle transfer by 3D fluorescence microscopy (I). The boxed areas in H and I are magnified in H1 and I1, respectively. (J and K) Proposed model for TNT-mediated organelle transfer. A cell forms actin-driven protrusions directed toward a target cell [(J), arrowhead]. TNT formation results in membrane continuity between connected cells. Organelles are unidirectionally transferred (arrow) via actin-mediated mechanisms. Red, F-actin; green, organelles. Scale bars, 20 ␮m.

pressed green fluorescent protein nor the small dye molecule calcein (fig. S7) were found to be transferred in detectable amounts between TNTconnected cells. Thus, with the exception of actin as a major structural component of TNTs, the small inner diameter of the stretched-membrane tubes, filled with F-actin, appears largely

to impede the passive transfer of soluble cytoplasmic molecules. To test whether plasmamembrane components could be transferred between TNT-connected cells, we analyzed the transfer of EGFP fused to the farnesylation signal of c-Ha-Ras (f-EGFP), a fusion protein that associated tightly with the membrane and spe-

cifically localized to the plasma membrane (fig. S8). f-EGFP was detected as discrete signals at the plasma membrane of those cells of population 2 that were connected via a TNT to cells of population 1 (Fig. 3, N to Q2). As in the fEGFP–expressing cells, transferred f-EGFP was found exclusively at the plasma membrane and displayed a partly continuous surface labeling, as well as patchy or raft-like signals (Fig. 3, N to Q2, arrowheads). Thus, plasma-membrane components can be indeed transferred between TNTconnected cells. To get more insight into the transfer of EGFP fusion proteins, we analyzed living cells of mixed populations by fluorescence videomicroscopy (1). After 24 to 48 hours, a weak fluorescence of f-EGFP was detectable in TNT-connected cells of population 2 (Fig. 4, A to C). This staining was in part continuous with the TNT labeling and covered large surface areas of the cells (fig. S9). To monitor the transfer of f-EGFP directly, we enhanced its transition by increasing the osmotic pressure selectively in one cell of the TNT-connected cell pair by microinjecting hyperosmotic solution (1). During the observation period of 10 min, this led to a continuous increase of f-EGFP fluorescence at the plasma membrane of the connected cell (Fig. 4, D to E2; see also fig. S10). Thus, plasma-membrane components could flow selectively between TNTconnected cells, which suggests that their membranes are continuously connected. Because the EGFP fluorescence of transferred synaptophysin-EGFP was barely above the detection limit, we analyzed the TNT-based exchange of membrane containers by labeling PC12 cells with green fluorescent 1,1⬘dioctadecyl-3,3,3⬘,3⬘-tetramethylindocarbocyanine perchlorate (DiI) or red fluorescent 3,3⬘dioctadecyloxacarbocyanine perchlorate (DiO) (1). Both membrane-specific dyes, well retained in cells (8) and frequently used as longterm tracers (9), were observed to be efficiently endocytosed in PC12 cells and thus served as markers of the endosomal or lysosomal pathway. It was possible to detect TNTs displaying unidirectional transfer of fluorescent organelles (Fig. 4F). This enabled us to track the movement of distinct organelles; we saw partly overlapping trajectories (Fig. 4G). We could follow organelles entering a TNT on one side, organelles being transported along the TNT, and organelles exiting the TNT into the connected cell (Fig. 4G, tracks 1 to 4, respectively; for details, see fig. S11). During this unidirectional transfer, accumulation and/or depletion of organelles could not be observed. Furthermore, a continuous and rapid translocation of organelles could be detected at any given point along the TNT (fig. S11), which was consistent with the existence of a direct intercellular transfer mechanism based on membrane continuity. When we analyzed TNT-connected cell pairs of mixed cultures consisting of one cell of each population, it became apparent that in 74%

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REPORTS (⫾6.1%) of all cases, one cell (Fig. 4, H and H1) accumulated organelles fluorescing in the second color (Fig. 4, I and I1). With respect to the low percentage of 11.3 (⫾2.8%) of all cells in culture displaying transfer, this value suggests a strong correlation between organelle transfer and the existence of a TNT connection. The transfer of organelles first became visible 2 hours after coculturing and, at later time points, showed as distinct cells harboring mostly organelles fluorescing in both colors (fig. S12). This indicates fusion of red- and green-labeled structures, a result consistent with reports on early endosomal fusion (10). The intercellular, TNT-dependent transfer of labeled organelles was also detected by analyzing mixed cultures of DiI- and DiO-labeled NRK cells (fig. S13). For a pair of TNT-connected cells, only one cell displayed both colors, which suggested transfer in one direction only. A quantitative analysis by fluorescence-activated cell sorting (FACS) revealed that the increase in number of cells with mixed fluorescence correlated with the increase in number of TNTs between cells after plating (fig. S4A). Performing transfer experiments close to 0°C, conditions that block exo-, endo-, or phagocytotic events (11, 12), we still detected organelle transfer between TNT-connected cells (fig. S14). Thus, the observed transfer did not depend on conventional exo-, endo-, or phagocytotic events. Organelle exchange could be blocked in the presence of latrunculin-B (fig. S14E), which strongly supported the presence of an actin-based transfer mechanism. The observation that functional TNTs were also found in cell cultures of lineages other than neuroendocrine cells (figs. S1 and S13) raises the possibility that TNTs represent a general cellular phenomenon occurring in long-range cell-to-cell communication. The transfer of endosomerelated structures through TNTs is consistent with the finding that similar structures, termed argosomes, facilitate the intercellular spread of wingless morphogens (13). Argosomes are thought to be exchanged between cells via sequential exo- and endocytotic events (14). The transfer of melanosomes between melanocytes and keratinocytes represents another riddle of organelle exchange (15). It has been proposed that this transfer occurs by means of local membrane fusion or phagocytotic mechanisms (15). Our finding that cells can actively exchange small membrane carriers through membrane channels provides evidence for a new principle of cell-to-cell communication based on membrane continuity between TNT-connected cells (Fig. 4, J and K). Provided that TNTs are present in tissue, reconsideration of previous interpretations of intercellular communication may be necessary. In this respect, the concept of membrane continuity between animal cells may also facilitate cell-to-cell transport of, e.g., transcription factors or ribonucleoparticles, as has been documented for the plant kingdom (16, 17).

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References and Notes

1. Materials and methods are available as supporting material on Science Online. 2. F. A. Ramirez-Weber, T. B. Kornberg, Cell 97, 599 (1999). 3. A. Karlsson et al., Nature 409, 150 (2001). 4. M. J. Hannah, A. A. Schmidt, W. B. Huttner, Annu. Rev. Cell Dev. Biol. 15, 733 (1999). 5. V. Mermall, P. L. Post, M. S. Mooseker, Science 279, 527 (1998). 6. D. A. Schafer et al., J. Cell Biol. 143, 1919 (1998). 7. R. J. Pelham Jr., F. Chang, Nature Cell Biol. 3, 235 (2001). 8. M. G. Honig, R. I. Hume, Trends Neurosci. 12, 333 (1989). 9. D. P. Kuffler, J. Comp. Neurol. 302, 729 (1990). 10. J. P. Gorvel, P. Chavrier, M. Zerial, J. Gruenberg, Cell 64, 915 (1991). 11. I. H. Pastan, M. C. Willingham, Science 214, 504 (1981). 12. M. Desjardins, L. A. Huber, R. G. Parton, G. Griffiths, J. Cell Biol. 124, 677 (1994). 13. V. Greco, M. Hannus, S. Eaton, Cell 106, 633 (2001). 14. K. Denzer, M. J. Kleijmeer, H. F. Heijnen, W. Stoorvogel, H. J. Geuze, J. Cell Sci. 113, 3365 (2000).

15. G. Scott, S. Leopardi, S. Printup, B. C. Madden, J. Cell Sci. 115, 1441 (2002). 16. K. Nakajima, G. Sena, T. Nawy, P. N. Benfey, Nature 413, 307 (2001). 17. W. Lucas, B.-C. Yoo, F. Kragler, Nature Rev. Mol. Cell Biol. 2, 849 (2001). 18. We thank A. Hellwig for generous help in electron microscopy, A. Kehlenbach and B. Schwappach for FACS analysis, J. Hammer for providing Dil2 antibody, A. Matus for providing EGFP-actin, R. Leube for providing synaptophysin-EGFP, and W. Franke and J. Leichtle for valuable comments on the manuscript. I.M. was supported by the Coimbra Group Hospitality Scheme and A.R. by the Landesgraduiertenstipendium Baden-Wu¨rttemberg; H.-H.G. was a recipient of grants from the Deutsche Forschungsgemeinschaft (SFB 488/B2, GE 550/3-2). Supporting Online Material www.sciencemag.org/cgi/content/full/303/5660/1007/ DC1 Materials and Methods Figs. S1 to S14 Movies S1 to S8 References 30 October 2003; accepted 10 December 2003

Direct Activation of Bax by p53 Mediates Mitochondrial Membrane Permeabilization and Apoptosis Jerry E. Chipuk,1 Tomomi Kuwana,1 Lisa Bouchier-Hayes,1 Nathalie M. Droin,1 Donald D. Newmeyer,1 Martin Schuler,2 Douglas R. Green1* The tumor suppressor p53 exerts its anti-neoplastic activity primarily through the induction of apoptosis. We found that cytosolic localization of endogenous wild-type or trans-activation–deficient p53 was necessary and sufficient for apoptosis. p53 directly activated the proapoptotic Bcl-2 protein Bax in the absence of other proteins to permeabilize mitochondria and engage the apoptotic program. p53 also released both proapoptotic multidomain proteins and BH3-only proteins [Proapoptotic Bcl-2 family proteins that share only the third Bcl-2 homology domain (BH3)] that were sequestered by Bcl-xL. The transcription-independent activation of Bax by p53 occurred with similar kinetics and concentrations to those produced by activated Bid. We propose that when p53 accumulates in the cytosol, it can function analogously to the BH3-only subset of proapoptotic Bcl-2 proteins to activate Bax and trigger apoptosis. The induction of apoptosis is central to the tumor-suppressive activity of p53 (1). Upon activation by DNA damage–induced or oncogene-induced signaling pathways, p53 promotes the expression of a number of genes that are involved in apoptosis, including those encoding death receptors (2, 3) and proapoptotic members of the Bcl-2 family (4, 5). In most cases, p53-induced apoptosis proceeds through mitochondrial release 1 Division of Cellular Immunology, La Jolla Institute for Allergy and Immunology, 10355 Science Center Drive, San Diego, CA 92121, USA. 2Department of Medicine III, Johannes Gutenberg University, D-55101 Mainz, Germany.

*To whom correspondence should be addressed. Email: [email protected]

of cytochrome c, which leads to caspase activation (6). Although most of the effects of p53 are ascribed to its function as a transcription factor, reports have suggested that the protein can also induce apoptosis independently of new protein synthesis (7–10). However, these studies have relied on ectopic expression of p53 or overexpression of mutants that lack transcriptional activity. Transcription-independent induction of apoptosis by p53 requires Bax and involves cytochrome c release and caspase activation, all of which occur in the absence of a nucleus, suggesting that p53 has the capacity to engage the apoptotic program directly from the cytoplasm (11). We therefore tested if endogenous p53 can engage the apoptotic program directly

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