Absolute Arrangement of Subunits in Cytoskeletal Septin Filaments in

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Absolute Arrangement of Subunits in Cytoskeletal Septin Filaments in Cells Measured by Fluorescence Microscopy Charlotte Kaplan,† Bo Jing,† Christian M. Winterflood,† Andrew A. Bridges,‡ Patricia Occhipinti,‡ Jürgen Schmied,§ Sören Grinhagens,∥ Thomas Gronemeyer,∥ Philip Tinnefeld,§ Amy S. Gladfelter,‡ Jonas Ries,† and Helge Ewers*,† †

Institute of Biochemistry, ETH Zurich, 8093 Zurich, Switzerland Life Science Center, Dartmouth College, Hanover, New Hampshire 03755, United States § Institute for Physical & Theoretical Chemistry, Braunschweig University, Hans-Sommer-Str. 10, 38106 Braunschweig, Germany ∥ Institute of Molecular Genetics and Cell Biology, University of Ulm, 89081 Ulm, Germany ‡

S Supporting Information *

ABSTRACT: We resolved the organization of subunits in cytoskeletal polymers in cells by light microscopy. Septin GTPases form linear complexes of about 32 nm length that polymerize into filaments. We visualized both termini of septin complexes by single molecule microscopy in vitro. Complexes appeared as 32 nm spaced localization pairs, and filaments appeared as stretches of equidistant localizations. Cellular septins were resolved as localization pairs and thin stretches of equidistant localizations. KEYWORDS: Septin, cytoskeleton, superresolution microscopy, single molecule, DNA origami he continuing improvement of superresolution fluorescence microscopy methods based on the sequential stochastic localization of individual molecules1,2 has opened the door to the investigation of the nanoscale architecture of multiprotein complexes in cells.3−5 While in vitro, distances down to few nanometers have been measured,6−8 in cells so far it has been impossible to resolve such distances. However, in a highly repetitive structure such as a cytoskeletal polymer, a localization precision in the range of 10 nm may suffice to resolve the relative positions of the monomers within the higher-order structure if they were of 20−30 nm length. The septins are a family of GTP-binding proteins with conserved function in the cell cycle.9−11 Four yeast septins assemble into rod-like complexes of about 32 nm length12 as palindromic hetero-octamers in a 2:2:2:2 stoichiometry.13 In vitro, these complexes polymerize further into pairwise aligned filaments that form bundles and rings.13−15 Septin function is best understood in yeast, where they form filamentous structures at the neck of dividing cells,16−18 which seem to be similar to filaments assembled in vitro from purified septin rods.17,19 However, the precise arrangement and organization of complexes in cellular filaments remains ambiguous. Here we set out to investigate whether septin complexes and their assembly order within filaments can be directly observed by single molecule localization microscopy in cells and whether the complexes assemble in an end-to-end fashion in cells. We labeled both termini of the rod-shaped septin complex and resolved the organization of the complexes in cells from the

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pattern emerging in single molecule localization microscopy in vitro and in cells. In purified septin complexes, we resolved the septin Cdc11, which occupies both ends of the complex as pairs of dots with a spacing of around 30−40 nm, consistent with the length of septin complexes. We show that filamentous structures assembled in vitro exhibit equidistant chains of localizations when we labeled Cdc11 and when we labeled Cdc10, the septin located in a pair at the center of the complex. In the filamentous fungus Ashbya gossypii, a well-studied modelsystem for septin assembly,15,20,21 we find that Cdc11 staining exhibits ∼35 nm spaced pairs of localizations and shows equidistant patterns in very thin filaments at around 35 nm distances as well, consistent with end-to-end assembly. Our results provide insight into the assembly of cytoskeletal filaments in cells at molecular resolution by fluorescence microscopy. The mirror-symmetric, 32 nm long yeast septin complex contains two Cdc11 molecules at its ends13 (Figure 1a) and is known to assemble end-to-end into pairwise aligned filaments in vitro. If Cdc11-molecules were labeled with an organic dye, the junctions between aligned pairs of complexes would thus be studded with four fluorophores (see Figure 1a). Such an arrangement greatly increases the chances of labeling and detection of each junction and thereby the resolution of the Received: February 19, 2015 Revised: April 30, 2015

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DOI: 10.1021/acs.nanolett.5b00693 Nano Lett. XXXX, XXX, XXX−XXX

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Figure 1. Detection of purified septin complexes as pairs of localizations. (a) Illustration of the experimental setup. Septin complexes were labeled at the terminal Cdc11 molecules, which are ∼32−35 nm apart with an organic dye. These dyes were then detected by single molecule localization microscopy experiments in vitro or in cells. (b) DNA-origami labeled at a spacing of 94 nm. (c) DNA-origami labeled at a spacing of 33 nm. (d) Negative stain transmission electron microscopy images of purified monodisperse S. cerevisiae septin complexes. (e) S. cerevisiae septin complexes labeled at Cdc11-GFP. Shown are for the different samples: single molecule superresolution microscopy reconstructed images (leftmost), close-ups of individual localization pairs (middle left), the overall distribution of all distances measured (middle right), and the distribution of average distances between localization pairs (right). Scale bars are 50 nm.

with Alexa Fluor 647 (AF647) and imaged them in a custombuilt single molecule microscopy setup23 under conditions favorable for single molecule localization microscopy, we detected pairs of point clouds of consistent spacing (Figure 1b). The point clouds resulted from the multiple localizations generated by each fluorophore in single molecule localization microscopy. When we plotted the distances between all individual localizations within 250 × 250 nm fields around these point clouds, the point clouds gave rise to a peak between 10 and 20 nm. We will refer to this peak as the “single molecule peak” as it represents the distances between multiple localizations of single dyes. In addition to this single molecule peak, we also detected a second peak of interlocalization distances at 90−100 nm (Figure 1b) resulting from the spacing between point clouds. When we then rendered individual localizations as Gaussian peaks and generated a cumulative image, we measured an average distance of 87.9 ± 9.0 nm (n = 100, N = 3) between these “intermolecule” peaks, consistent with the distance between the fluorophores on the DNA origami. We concluded that we can accurately measure the distance between individual fluorophores at ∼90 nm spacing. Next we labeled DNA-origami structures at a spacing of around 33 nm, similar to the length of septin complexes (Figure 1c). In this measurement, the single molecule peak was very close to the intermolecule peak, and we measured a

spacing between the termini of adjacent monomers in the filament. To investigate whether such a measurement would be technically feasible, we simulated a septin filament assembled from pairwise aligned, 32 nm long complexes (see Supporting Information, Figure 1a) and generated in silico superresolution images according to a number of variables, such as the labeling rate, the average localization precision, the distribution around the average localization precision and the number of background localizations per area. As an alternative model, we simulated septin complexes assembled side-by-side into filaments (see Supporting Information, Figure 1b) to ask under which labeling and imaging conditions alternative models of septin organization could be distinguished. From our simulations we found that these simple models could be easily distinguished when we reduced the detection efficiency to 50% and allowed for a distribution of localization precisions around 10 nm with a σ of 5 nm, suggesting that end-to-end assembly of septin complexes into pairwise filaments is robustly detectable using single molecule localization microscopy. We first aimed to test our imaging system and analysis approach with a simplified system, where individual dyes are spaced at a precisely determined distance. To do so, we made use of DNA-origami structures that were labeled at defined positions with dyes22 (see Supporting Information, Figure 2). When we labeled these structures in vitro at a spacing of 94 nm B

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Figure 2. Detection of end-to-end assembly of septin complexes in vitro. (a) Model of a septin complex labeled on Cdc11 (left). When assembled into filaments, such complexes will yield equidistant localizations at 32 nm spacing (right). (b) Fluorescence micrograph of in vitro assembled Cdc11-GFP labeled septin complexes (left). Superresolution image of the same field of view (right). The scale bar is 200 nm. Gaussian reconstructed single molecule localization images of Cdc11-labeled septin complex assemblies. The scale bar is 100 nm. (c) Model of a septin complex labeled on Cdc10 (left). Individually, such complexes will be detectable only as dots, but when assembled into filaments, such complexes will yield equidistant localizations at 32 nm spacing. (d) Gaussian reconstructed single molecule localization images of Cdc10-labeled septin complex assemblies. The scale bar is 100 nm. (e) Average spacing between localization along filament axis for Cdc10-SNAPf and Cdc11-GFP. (f) Average length of Cdc10-SNAPf and Cdc11-GFP filaments. (g) Average thickness of filaments. (e−g) Boxes are in the 25th to 75th percentile of measured values; the central line represents the median, and the whisker line represents the range of all measured values.

distance between cumulative Gaussian peaks of 31.7 ± 4.1 nm (n = 90, N = 3). These measurements show that we can detect distances in the range of the size of septin complexes in this simplified system. We then purified recombinant Saccharomyces cerevisiae septin complexes in high salt (300 mM NaCl), a condition under which septin complexes do not polymerize in vitro24 (Figure 1d, see also Supporting Information, Figure 3a). In these complexes, we tagged Cdc11, the septin occupying both termini of the rod, with GFP.25 When we added Cdc11-GFP complexes to coverglass with a functionalized and passivated surface, labeled them with AF647 coupled anti-GFP nanobodies23 and performed single molecule localization microscopy, we could detect pairs of localizations with a spacing of 38.6 ± 8.2 nm (n = 99, N = 3, Figure 1e), consistent with the size of septin complexes measured in electron microscopy.13,24 Such pairs were not detectable above background when the experiment was performed with GFP alone (data not shown). We also occasionally detected linear equidistant triplets of localizations, suggesting that two septin complexes were attached to each other in an end-to-end fashion. In quality

control experiments in electron microcopy, we likewise occasionally found complexes twice as long as a single rod (see Supporting Material, Figure 3b,c). Assuming that not all Cdc11 molecules likely are labeled, this observation explains the existence of a second peak in the distance plot for septin complexes at around 70 nm (Figure 1e), consistent with the length of two adjacent complexes. We concluded that the absolute position and orientation of individual septin complexes can be measured by single molecule localization microscopy in vitro. We next aimed to detect individual septin complexes when assembled into filaments. We expected septin complexes to assemble end-to-end into linear filaments, resulting in a chain of localizations with a spacing of about 32 nm (Figure 2a). To promote filament assembly, we diluted purified Cdc11-GFP containing septin complexes into low salt buffer to 12 nM, a condition known to induce septin complex polymerization.13 We then incubated them for 60 min in solution together with AF647 coupled anti-GFP nanobodies before adding them to polylysine-coated coverglass. We observed filamentous structures of 1−5 μm length and many smaller structures in the C

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Cdc11 molecule was tagged and accessible for labeling and then labeled the cells with BG-AF647. This resulted in bright staining of elongated filamentous septin structures and some diffusely located Cdc11-SNAPf staining as observed in TIRF imaging (Figure 3a).

diffraction limited GFP channel. When we imaged these structures in single molecule localization microscopy, many of the longer filaments appeared as very thin, straight chains of periodic localizations. These filaments often were straight over more than 1 μm at a thickness of 30 nm or less, while some filaments were bent and some even had kinks that were not apparent in the diffraction-limited image. A periodicity of localizations at ∼32 nm was especially apparent in shorter structures in which only 3−10 septin complexes seemed to be assembled (Figure 2b). Longer filaments often seemed to be bundled as judged from their thickness (up to ∼100 nm) and clear patterns of localizations could not be detected, likely because filaments were not aligned in bundles (see Supporting Information, Figure 6b,c). These bundles could be curved as well up to a degree consistent with the formation of 500−600 nm rings around yeast bud-necks. To verify that the periodicity we observed was indeed due to end-to-end assembly of 35 nm long complexes, we also used a different labeling approach. We purified septin complexes in which the septin in the center of the octameric complex, Cdc10, was tagged with a SNAPf-tag,26,27 and subsequently labeled in the test tube with AF647-benzylguanine. We reasoned that if septin complexes assembled end-to-end into filaments, we would expect Cdc10-labeled complexes to show the same 32 nm spacing as Cdc11-labeled complexes (Figure 2c). When we incubated Cdc10-labeled complexes as described before for 60 min in low salt buffer and added this sample to coverglass, we could indeed observe a similar spacing in filaments (Figure 2d). The average distance between Cdc11 or Cdc10 molecules in such filaments was 33.6 ± 14.7 nm (n = 182, N = 2) for Cdc10 and 31.4 ± 11.3 nm (n = 140, N = 2) for Cdc11 (Figure 2e) consistent with the distance measured for individual septin complexes in electron microscopy.12,13 Filaments were on average 1.53 ± 0.66 μm (Cdc10, n = 22) and 1.26 ± 0.77 μm (Cdc11, n = 24) long (Figure 2f) consistent with previous observations.25,26,28 When we measured the average thickness of single filaments exhibiting clearly spaced chains of localizations as full width halfmaximum of the Gaussian rendering of the cumulative localizations, we found an average thickness of 30.7 ± 9.2 nm (Cdc10, n = 22) and 37.6 ± 7.4 nm (Cdc11, n = 24) (Figure 2g). This thickness is in the range observed for microtubules in single molecule localization microscopy and consistent both with single and pairwise-aligned filaments and it may even accommodate more than two aligned filaments. Since we observed consistent spacing, it is unlikely that such structures represent unaligned bundles. Thus, we can detect the relative positions of septin complexes in end-to-end assembled filaments by single molecule localization microscopy. We next aimed to determine whether septin complexes assemble end-to-end into filaments in cells as well. The globular shape of S. cerevisiae however is not amenable to resolve the individual positions of molecules in three dimensions due to the lower resolution in 3D localization microscopy. We thus made use of the filamentous fungus A. gossypii, in which septins also assemble into 30−35 nm hetero-octamers and form elongated filaments in vitro.15 In contrast to S. cerevisiae, A. gossypii cells have a tubular shape, and membrane-associated septin structures are thus aligned with the coverglass and readily accessible in total internal reflection fluorescence (TIRF) illumination for single molecule microscopy. We generated an Ashbya strain in which Cdc11 was endogenously tagged with a SNAPf-tag to ensure that every

Figure 3. Detection of septin complexes in cells. (a) Total internal reflection fluorescence microscopy image of an A. gossypii cell expressing Cdc11-SNAPf and labeled with BG-AF647. Shown is a hypha exhibiting some septin filaments and diffuse septin staining at its tip. The scale bar is 1 μm. (b) Single molecule localization microscopy image of the inset in panel a. Several pairs of localizations with 30−40 nm spacing are emphasized by boxes. Scale bar is 200 nm. (c1, c2) Single molecule localization microscopy reconstructed images depicting areas from (b) with several localizations arranged in 30− 40 nm spacing in close proximity, some of which are aligned end-toend. Scale bars are 50 nm. (d) Distribution of pairwise distances between localizations in (b). Arrows point to peaks around 30−35 nm, 70−80 nm, and 100−110 nm.

When we performed single molecule localization microscopy in these cells, we found different modes of organization. In areas where septin staining was scarce (Figure 3a), we found many individual, dot-like localizations frequently arranged in pairs (Figure 3b) or small assemblies (Figure 3c) with spacings of around 32 nm. In fact, when we plotted the interlocalization distances in a region of interest such as Figure 3b, we could detect a periodicity of peaks at ∼30−35 nm, 70−80 nm, and 100−110 nm consistent with one, two, and three complex lengths (Figure 3d), suggesting that a substantial number of complexes were present in straight, dimeric, or trimeric end-to-end assemblies (see also Figure 3c1). This was not necessarily expected as the density of septin signal in cells may overcome any detectable peaks. Interestingly we also observed a peak at ∼50 nm, which may represent the diagonal distance found between parallel complexes (see Figure 3c2). In other areas of the cells, as in vitro, we found linearly aligned stretches of localizations in structures of varying length and thickness (Figure 4a). It seemed that individual filaments were occasionally associated laterally in the plane of the D

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microscopy studies found septins to assemble into thin, continuous filamentous structures.17 Our findings demonstrate that these filaments likely represent chains of full-length complexes with regularly spaced Cdc11 molecules. In vitro, we found regular, ∼32 nm equidistant localizations both when we labeled Cdc11 and when we labeled Cdc10. This observation can only be explained by a model of the septin complex where Cdc11 occupies the termini of the complex and Cdc10 is located to the center of the complex as proposed by Bertin et al.13 It is also not consistent with any arrangement besides end-to-end assembly of septin complexes into filaments. The filaments were around 30 nm thick, which could be explained by single filaments as well as by pairwise aligned filaments. If a collapse of the coiled coils as proposed by Thorner et al.,30 is assumed, even four to six closely aligned filaments could be explained by our findings. Given the resolution precision of our experiments, it is not possible to distinguish between single and pairwise-aligned filaments. It is clear, however that all detected septin molecules must be in register as otherwise the intermolecular spacing would not be detectable. When we analyzed thicker structures, we could indeed not detect the regular spacing of complexes. Such structures likely represented bundles of unaligned filaments. Thus, depending on the size of septin bundles, septin filaments can be seen either in exact register or in less organized assemblies. Based on the observation of pairwise aligned filaments of purified yeast septin complexes in electron microscopy and the detection of electron dense filamentous structures that contain septins in intact yeast cells it has been postulated that septin complexes assemble end-to-end into filaments in cells as well. However, cellular septins interact with a number of molecules and membranes and may well appear filamentous because they associate with other structures rather than polymerizing into continuous filaments. The filaments we observed in cells strikingly resembled the filaments found in vitro both in terms of spacing between localizations as well as thickness. The structures we observed in cells contained relatively long, straight and very thin strings of localizations, likely from a single or two pairwise aligned filaments. A recent electron tomography study showed that yeast septin complexes can form more complex structures than pairwise filaments in vitro when incubated with Cdc42 and Gic1.28 The authors demonstrated that aligned septin filaments are grouped into cables by a Cdc42-Gic1 complex. These cables are 20−30 nm thick, which is in agreement with what we find in cells. The structures we observe may thus be organized in such cables. In conclusion, we find that septin complexes can assemble end-to-end in cells, providing definitive information about the cellular organization of a conserved, essential filament-forming protein family. Our imaging approach opens the door to a detailed investigation of the relative organization of septins and septin interacting molecules in different cellular processes. Such experiments will give important insight into previously inaccessible aspects of septin function such as their role as molecular scaffolds and the organization of diffusion barriers. Besides septin biology, our work provides a stepping-stone for future investigations of how the function of multiprotein complexes is encoded in the organization of their constituting molecules.

Figure 4. Septin complexes assemble end-to-end in cells. (a) Total internal reflection fluorescence microscopy image of an A. gossypii cell expressing Cdc11-SNAPf and labeled with BG-AF647. Septin filaments aligned with the cell axis are prominently labeled at the bottom surface of the cell. The scale bar is 500 nm. (b) Single molecule localization microscopy of an area of the same cell shows discrete localizations in filaments and filament bundles in regular patterns. The scale bar is 500 nm. (c) Detail of a septin structure from (b) exhibiting clearly resolved repetitive ∼32 nm distances. The area in the outer broken line is 65 × 650 nm. The area in the inner broken line is shown in (d). The scale bar is 100 nm. (d) Reconstructed image of a part of the filament in (b) after rejection of all localizations with localization precision ≤10 nm. The filament axis is emphasized by a white line. Below at the same scale the intensity plot along the filament axis of the Gaussian rendered reconstruction of all localizations (broken line) or only localizations with a precision ≤10 nm (solid line) is shown. Above the graph the distances between the peaks in nanometers are shown.

membrane as we often observed parallel chains of localizations that appeared as filaments. Such filaments were on average 1.15 ± 0.40 μm long (n = 21, N = 5) and varied greatly in thickness (see Figure 4b). When we analyzed these filaments more closely, we found individual filaments that demonstrated a spacing of individual localizations of 35.9 ± 4.0 nm (n = 23, N = 4, Figure 4c). When we rejected localizations that could not be localized with at least 10 nm precision in such filaments in a “lucky imaging” approach,29 the spacing of 30−40 nm became clearly apparent (Figure 4d). Such filaments exhibited a full width half-maximum of around 20 nm at length of up to 30 to 50 times more than this, consistent with very thin, straight filaments, likely single or pairwise aligned as found in vitro. When we compared such cellular filaments to our simulations they likened pairwise aligned end-to-end filaments, but not side-by-side assembly of septin complexes (see Supporting Information, Figure 4). In fact, we never found any arrangement resembling side-by-side assembly in cells or in vitro. We concluded that septin complexes assemble end-to-end in cells. When we investigated assemblies formed from purified septin complexes by single molecule localization microscopy, we consistently found linear stretches of regularly spaced localizations with around 30−35 nm gaps. Previous electron E

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(2) Rust, M. J.; Bates, M.; Zhuang, X. Nat. Methods 2006, 3, 793− 795. (3) Szymborska, A.; de Marco, A.; Daigle, N.; Cordes, V. C.; Briggs, J. A. G.; Ellenberg, J. Science 2013, 341, 655−658. (4) Xu, K.; Zhong, G.; Zhuang, X. Science 2012, 339, 452−456. (5) Kanchanawong, P.; Shtengel, G.; Pasapera, A. M.; Ramko, E. B.; Davidson, M. W.; Hess, H. F.; Waterman, C. M. Nature 2010, 468, 580−584. (6) Raab, M.; Schmied, J. J.; Jusuk, I.; Forthmann, C.; Tinnefeld, P. ChemPhysChem 2014, 15, 2431−2435. (7) Jungmann, R.; Avendaño, M. S.; Woehrstein, J. B.; Dai, M.; Shih, W. M.; Yin, P. Nat. Methods 2014, 11, 313−318. (8) Vaughan, J. C.; Jia, S.; Zhuang, X. Nat. Methods 2012, 9, 1181− 1184. (9) Hartwell, L. H. Exp. Cell Res. 1971, 69, 265−276. (10) Kinoshita, M.; Kumar, S.; Mizoguchi, A.; Ide, C.; Kinoshita, A.; Haraguchi, T.; Hiraoka, Y.; Noda, M. Genes Dev. 1997, 11, 1535− 1547. (11) Surka, M. C.; Tsang, C. W.; Trimble, W. S. Mol. Biol. Cell 2002, 13, 3532−3545. (12) Field, C. M.; al-Awar, O.; Rosenblatt, J.; Wong, M. L.; Alberts, B.; Mitchison, T. J. J. Cell Biol. 1996, 133, 605−616. (13) Bertin, A.; McMurray, M. A.; Grob, P.; Park, S.-S.; Garcia, G.; Patanwala, I.; Ng, H.-L.; Alber, T.; Thorner, J.; Nogales, E. Proc. Natl. Acad. Sci. U. S. A. 2008, 105, 8274−8279. (14) Garcia, G.; Bertin, A.; Li, Z.; Song, Y.; McMurray, M. A.; Thorner, J.; Nogales, E. J. Cell Biol. 2011, 195, 993−1004. (15) Meseroll, R. A.; Howard, L.; Gladfelter, A. S. Mol. Biol. Cell 2012, 23, 3391−3406. (16) Byers, B.; Goetsch, L. J. Cell Biol. 1976, 69, 717−721. (17) Ong, K.; Wloka, C.; Okada, S.; Svitkina, T.; Bi, E. Nat. Commun. 2014, 5, 5698. (18) Bertin, A.; McMurray, M. A.; Pierson, J.; Thai, L.; McDonald, K. L.; Zehr, E. A.; Garcia, G.; Peters, P.; Thorner, J.; Nogales, E. Mol. Biol. Cell 2011, 23, 423−432. (19) McMurray, M. A.; Bertin, A.; Garcia, G.; Lam, L.; Nogales, E.; Thorner, J. Dev. Cell 2011, 20, 540−549. (20) DeMay, B. S.; Bai, X.; Howard, L.; Occhipinti, P.; Meseroll, R. A.; Spiliotis, E. T.; Oldenbourg, R.; Gladfelter, A. S. J. Cell Biol. 2011, 193, 1065−1081. (21) Meseroll, R. A.; Occhipinti, P.; Gladfelter, A. S. Eukaryotic Cell 2013, 12, 182−193. (22) Schmied, J. J.; Gietl, A.; Holzmeister, P.; Forthmann, C.; Steinhauer, C.; Dammeyer, T.; Tinnefeld, P. Nat. Methods 2012, 9, 1133−1134. (23) Ries, J.; Kaplan, C.; Platonova, E.; Eghlidi, H.; Ewers, H. Nat. Methods 2012, 9, 582−584. (24) Frazier, J. A.; Wong, M. L.; Longtine, M. S.; Pringle, J. R.; Mann, M.; Mitchison, T. J.; Field, C. J. Cell Biol. 1998, 143, 737−749. (25) Bridges, A. A.; Zhang, H.; Mehta, S. B.; Occhipinti, P.; Tani, T.; Gladfelter, A. S. Proc. Natl. Acad. Sci. U.S.A. 2014, 111, 2146−2151. (26) Renz, C.; Johnsson, N.; Gronemeyer, T. BMC Biotechnol. 2013, 13, 60. (27) Gautier, A.; Juillerat, A.; Heinis, C.; Corrêa, I. R.; Kindermann, M.; Beaufils, F.; Johnsson, K. Chem. Biol. 2008, 15, 128−136. (28) Sadian, Y.; Gatsogiannis, C.; Patasi, C.; Hofnagel, O.; Goody, R. S.; Farkasovsky, M.; Raunser, S. Elife 2013, 2, e01085. (29) Cronin, B.; de Wet, B.; Wallace, M. I. Biophys. J. 2009, 96, 2912−2917. (30) Bertin, A.; McMurray, M. A.; Thai, L.; Garcia, G.; Votin, V.; Grob, P.; Allyn, T.; Thorner, J.; Nogales, E. J. Mol. Biol. 2010, 404, 711−731.

ASSOCIATED CONTENT

S Supporting Information *

Materials and methods, Figures 1−6. The Supporting Information is available free of charge on the ACS Publications website at DOI: 10.1021/acs.nanolett.5b00693.

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AUTHOR INFORMATION

Corresponding Author

*E-mail [email protected]. Present Addresses

C.K.: Department of Molecular and Cell Biology, University of California Berkeley, Berkeley, CA 94720-3202, USA. B.J.: Biological Physics Research Group, Clarendon Laboratory, Department of Physics, University of Oxford, Oxford OX1 3PU, United Kingdom. C.M.W. and H.E.: Randall Division of Cell and Molecular Biophysics, King’s College London, SE1 1UL, United Kingdom. J.R.: Cell Biology and Biophysics Unit, EMBL Heidelberg, Meyerhofstrasse 1, 69117 Heidelberg, Germany. H.E.: Freie Universität Berlin, Institute of Chemistry and Biochemistry, Thielallee 63, 14195 Berlin, Germany. Author Contributions

H.E. conceived the project; C.K. and H.E. designed experiments; C.K. performed experiments; B.J. provided analytical tools; H.E., C.K., C.W., and J.R. analyzed data; A.G., T.G., L.O., S.G., and A.B. provided purified septin complexes; P.O. cloned A. gossypii strain; J.S. and P.T. provided DNA origami; H.E. and C.K. wrote the manuscript. Funding

The authors acknowledge support from the NCCR Neural Plasticity and Repair, the Holcim Foundation, the Swiss National Fund, an EMBO short-term fellowship (CK), a Boehringer Ingelheim Travel grant (CK), a Marie Curie Fellowship (JR, CMW) and the NCCBI. J.J.S. and P.T. were supported by the Biophotonics IV program of the Federal Ministry of Education and Research (BMBF, VDI) (13N11461). P.O., A.A.B., and A.S.G were supported by funding from the National Science Foundation (MCB-507511) and the National Institutes of Health (GM100160). Notes

The authors declare no competing financial interest.

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ACKNOWLEDGMENTS The authors thank Ingmar Schoen for help with origami labeling and data analysis, Markus Mund for help with the setup in the Ries lab and all members of the Ewers and Ries laboratories for helpful discussions. The authors thank Gleb Shtengel for help with the purchase of SNAP-Surface® AF647. We thank the EMBL electron microscopy facility, especially Rachel Mellwig, for help with electron microscopy. The authors thank Enrico Pibiri for AFM imaging of the DNA-origami structures.

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ABBREVIATIONS BG, benzylguanine; AF, Alexa Fluor; GFP, green fluorescent protein REFERENCES

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DOI: 10.1021/acs.nanolett.5b00693 Nano Lett. XXXX, XXX, XXX−XXX