Super-resolution Microscopy of Clickable Amino Acids Reveals the

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Super-resolution Microscopy of Clickable Amino Acids Reveals the Effects of Fluorescent Protein Tagging on Protein Assemblies )

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Ingrid C. Vreja,†,‡ Ivana Nikic,§ Fabian Go¨ttfert, Mark Bates, Katharina Kro¨hnert,† Tiago F. Outeiro,^ Stefan W. Hell, Edward A. Lemke,§ and Silvio O. Rizzoli*,† †

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Department of Neuro- and Sensory Physiology and ^Department of NeuroDegeneration and Restorative Research, University Medical Center Göttingen, 37073 Göttingen, Germany, ‡International Max Planck Research School for Molecular Biology, Göttingen, Germany, §Structural and Computational Biology Unit, EMBL, 69117 Heidelberg, Germany, and Department of Nanobiophotonics, Max Planck Institute for Biophysical Chemistry, 37077 Göttingen, Germany

ABSTRACT The advent of super-resolution microscopy (nanoscopy)

has set high standards for fluorescence tagging. Fluorescent proteins (FPs) are convenient tags in conventional imaging, but their use in nanoscopy has been questioned due to their relatively large size and propensity to form multimers. Here, we compared the nanoscale organization of proteins with or without FP tags by introducing the unnatural amino acid propargyl-L-lysine (PRK) in 26 proteins known to form multimolecular arrangements and into their FP-tagged variants. We revealed the proteins by coupling synthetic fluorophores to PRK via click chemistry and visualized them using ground-state depletion microscopy followed by individual molecule return, as well as stimulated emission depletion microscopy. The arrangements formed by the FP-tagged and nontagged proteins were similar. Mild, but statistically significant differences were observed for only six proteins (23% of all proteins tested). This suggests that FP-based nanoscopy is generally reliable. Unnatural amino acids should be a reliable alternative for the few proteins that are sensitive to FP tagging. KEYWORDS: click chemistry . fluorescent protein . protein engineering . super-resolution microscopy . unnatural amino acid . GSDIM . STED

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he recent development of lens-based super-resolution fluorescence microscopy or nanoscopy1,2 has enabled the investigation of macromolecular complexes with a level of detail reaching beyond the size of many molecular complexes (10
30 nm).3,4 Exploring molecular-sized elements by fluorescence microscopy has become realistic. Yet limitations are increasingly set by the confidence with which the target molecules can be labeled. The labels used for fluorescence nanoscopy must not hinder the localization, function, and/or structure of their targets. This characteristic is important in all imaging techniques and becomes especially relevant in superresolution procedures. Since they were introduced before the advent of superresolution microscopy, many labels and labeling strategies fail in this attempt. Some VREJA ET AL.

labels, including antibodies, may place the fluorophores too far from the targets,5
7 yielding images that do not faithfully reproduce their spatial distributions. Antibodies may also induce the clustering of target proteins, especially when applied on live or on insufficiently fixed cells.8 Genetically encoded labels such as fluorescent proteins (FPs) or fluorophore-binding protein domains also need to be carefully tested. Their size of a few nanometers and their tendency to form multimeric arrangements may interfere with the localization of the target proteins. For example, FPs may induce protein clusters, which are occasionally evident in nanoscopy recordings.9 Nevertheless, the ease of use of FPs, especially in living cells, indicates that they will continue to be crucial for microscopy. Their use in nanoscopy techniques has been VOL. XXX

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* Address correspondence to [email protected]. Received for review July 17, 2015 and accepted October 24, 2015. Published online 10.1021/acsnano.5b04434 C XXXX American Chemical Society

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RESULTS AND DISCUSSION We set out to compare several proteins containing only the UAA tags with their FP-coupled chimeras (also VREJA ET AL.

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enhanced by the recent introduction of a nanobody for green fluorescent protein (GFP). Nanobodies coupled to chemical fluorophores optimized for any superresolution technique can now be used to reveal GFP chimeras.5,6 It is therefore important to test the effect of FPs on protein organization in general, by comparing a number of FP-tagged proteins with their nontagged counterparts. Antibody stainings and FP patterns are broadly similar for most proteins in diffraction-limited microscopy.10 But when differences are noted in nanoscopy,11 it is difficult to state whether the FP labeling or the antibody staining is at fault. To test the effects of the FP tag, it is necessary to study both the FP-tagged and nontagged proteins in parallel, with a nanoscopy label that is much smaller than FPs and thus avoids FP-related problems. A tool that would meet these conditions has been recently described: the genetic encoding of unnatural amino acids (UAAs) into specific proteins.12,13 This system relies on expressing appropriate pairs of tRNAs and aminoacyl-tRNA synthetases (tRNA/RS), along with a modified gene coding for the target protein, which contains an Amber stop codon (TAG). Orthogonal tRNA/RS pairs are typically used, which selectively recognize the Amber codon. This leads to the site-specific incorporation of the UAA of choice during translation14,15 The incorporation of the UAA is followed by coupling to fluorophores. For example, the UAA propagyl-L-lysine (PRK), which contains an alkyne group, is identified by fluorophores carrying an azide functionality, through copper-mediated azide
alkyne cycloaddition, also termed “click reaction”.14
16 The fluorophore is typically coupled to the UAA only after cell fixation. This implies that the proteins are allowed to behave freely, without interference from fluorescence labeling, until fixation at the desired time point. This strategy thus fulfills all the requirements of a tool for testing FPs, since the small UAA tag produces only minimal changes to the protein, but nevertheless allows the near-quantitative fluorescent labeling of the targets.14 We have used this system to test 26 proteins that are known to form various types of multimolecular arrangements. Tagging with GFP or YFP had no significant effects on the organization of 20 of the proteins. Relatively mild effects were seen on the remaining six proteins. This implies that FPs such as GFP or YFP are reliable in nanoscopy, although care must still be taken in their application. Other FPs that are more prone to multimerization should be tested especially carefully. To help with such efforts, we also provide a detailed protocol for comparing, within ∼10 days, the behavior of the GFP-tagged and nontagged versions of different proteins of interest.

Figure 1. Rationale of the experiments. Proteins without FP tags and their FP-tagged variants were subjected to PRK incorporation, followed by copper(I)-catalyzed click reactions with a synthetic fluorophore. The visualization of the latter enables the nanoscopy comparison between the molecular arrangements made by the FPtagged proteins and those of the original, nontagged variants.

coupled to the UAA tags; Figure 1). We relied on the wild-type tRNA/RS pair from Methanosarcina mazei, which can encode PRK in response to the Amber codon.14,15,17,18 We tested 11 transmembrane proteins, including the SNARE fusion proteins syntaxin 1, syntaxin 6, syntaxin 7, syntaxin 13, Vti1a-β, VAMP2 (synaptobrevin 2), VAMP4,19 a serotonin receptor (5HT1a),20 the insulin receptor (IR),21 and two synaptic vesicle proteins (synaptotagmin 1 and synaptophysin).22 We also tested six membrane-attached proteins: three SNAREs, which are permanently attached to the plasma membrane by palmitoylation (SNAP-25, SNAP-23, and SNAP-29),19 and three Rab proteins, which shuttle between endosomal or vesicular membranes and the cytosol (Rab3, Rab5, and Rab7).23 Finally, we tested nine soluble proteins, including proteins involved in endocytosis (amphiphysin, AP-2 μ), in exocytosis (complexin 1, Doc2R, Munc18-1, R-synuclein, synapsin Ia), in endosomal function (phosphatidylinositol-4-phosphate 5-kinase type I gamma, referred to as PIPKIγ), and in cytoskeletal function (β-actin).24,25 While a thorough discussion of these proteins is beyond the purpose of this work, we note that all have been shown to form, or participate in, protein domains, clusters, and other types of supramolecular arrangements. All of these proteins have been studied in the past by coupling them to FP tags: syntaxin 1;26 syntaxin 6, syntaxin 13, Vti1a-β, and VAMP2;27,28 synaptophysin and synaptotagmin; 11 VAMP4;29 R-synuclein; 30
32 Rab proteins;33,34 SNAP-25; 35 5HT1a; 36 insulin receptor (IR);37 synapsin;38 Munc18-1;39 complexin;40 syntaxin 7;41 amphiphysin;42 AP-2 μ;43 Doc2R;44 PIPKIγ;45 β-actin;46 SNAP-23;27 SNAP-29.34 VOL. XXX

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ARTICLE Figure 3. Fluorescent labeling of PRK-containing cells. To determine the specificity of the click chemistry reaction, we analyzed the intensity of the Star635P-azide fluorescence for cells expressing R-synuclein-GFP that were click labeled. (A) The fluorescence of the chemical dye is linearly correlated with the fluorescence of the GFP (i.e., with the expression of the UAA-containing protein), which argues that the reaction is specific. Symbols indicate averages of 172
1902 cells for R-synuclein from a single typical experiment. (B) The images show one transfected cell, surrounded by many nontransfected cells, whose nuclei are visible in the DAPI staining. Scale bar, 20 μm.

Figure 2. FP expression is specific in transfected cells. We have assessed the specificity of PRK incorporation for the 26 proteins included in this study. Control samples that were transfected, but were not subjected to PRK addition to the medium, are shown in the left panels (no UAA). No expression was detected under these conditions, indicating that the constructs strictly require the PRK incorporation. In the panels on the right we show representative epifluorescence images of cells that have been exposed to PRK, which enabled the expression of the proteins of interest, along with the respective FPs. Scale bar, 20 μm. VREJA ET AL.

We introduced Amber codons into all of the proteins, as well as into their FP-coupled chimeras (Figure S1, Tables S2 and S3). We used the most common FP tags, green and yellow fluorescent proteins, GFP and YFP (Figure S1). To test whether PRK was incorporated correctly, we monitored the expression of the FP-tagged constructs, in the presence or absence of PRK (Figure 2). As the FP tag was always placed in the protein sequence after the Amber codon, our experimental design predicts that no FP expression should be observed in cells cultured in the absence of PRK, as in these conditions the protein translation stops at the Amber codon. In contrast, when PRK is added to the medium, the Amber codon is no longer used as a stop, and the translation of the FP tag is enabled. This was indeed the case: no FP expression was observed in the absence of PRK, and its addition to the medium induced a substantial fraction of the cells to express the proteins (Figure 2). Having ascertained that the UAA incorporation functioned correctly, we next tested whether the click reaction was sufficiently accurate. The PRK side chains of the proteins were revealed, after fixation, by incubating the cells with azide-containing variants of the red fluorophores AlexaFluor647 or Star635P. We observed no Alexa or Star labeling in nontransfected cells or in cells grown in the absence of PRK. Figure 3 shows a transfected cell, in which the expressed GFP is visible (green), along with the chemical fluorophores that had been coupled to the proteins through the click reaction (red). No red labeling is discernible in the nontransfected cells (whose nuclei are revealed in the particular figure by DAPI staining). Moreover, in cells that did express the proteins of interest, the chemical fluorophore labeling correlated linearly with the amount of protein expressed, demonstrating that the click reaction was highly specific (Figure 3). VOL. XXX

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ARTICLE Figure 4. FP tags do not change the molecular arrangements of the majority of the investigated proteins. The figure shows representative images from ultrathin sections of melamin-embedded cells expressing nontagged proteins (no FP; left panels) and their chimeric counterparts (FP-tagged, right panels). All samples were allowed to incorporate PRK, were subsequently coupled to AlexaFluor647-azide, and were imaged using GSDIM microscopy. We would like to note that the AlexaFluor647 fluorescence is shown in all panels and not the FP fluorescence. Scale bar, 500 nm. The images were analyzed by fitting Gaussian functions onto the fluorescent spots (protein clusters), and bar plots were generated for the spot sizes and the peak intensities (mean ( SEM from, on average, ∼100 spots per protein). Data for FP-tagged and nontagged proteins are depicted in green and black, respectively. The Student's t test was used for assessing statistical significance (p < 0.05 *, p < 0.01 **, p < 0.001 ***). VREJA ET AL.

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For high-resolution imaging, we increased the imaging contrast by embedding the cells in plastic material (melamine) and by cutting them into ultrathin sections (see Materials and Methods for details). Protein assemblies were difficult to detect when investigated by conventional microscopy, but became evident when using ground-state depletion followed by individual molecule return (GSDIM) microscopy.47 The resolution of this technique, which uses the same principles and tools as stochastic optical reconstruction microscopy (STORM), reaches 20
30 nm. An overview of the results is shown in Figure 4. The use of ultrathin sections results in only a handful of molecules of interest in each field of view. While this removes the context of the cell and renders all of the images relatively similar, it does have the advantage that it also removes background and reduces drastically the complexity of the images. This enabled us to measure the size and the peak intensity for each of the fluorescent spots, restricting the analysis to multimolecular arrangements, whose intensity was beyond that of single molecules. The size of the FP-tagged protein assemblies changed significantly, when compared to the nontagged proteins, only for PIPKIγ (whose FP chimera generated larger and more intense clusters), VAMP4 (smaller and less intense clusters), and Vti1a-β (slightly larger clusters). In addition, the intensity of the protein clusters increased for β-actin and Munc18-1 and decreased for amphiphysin. VREJA ET AL.

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Figure 5. The FP tag does not change the protein organization in STED microscopy. Representative images of membrane sheets derived from cells expressing SNAP-25 (upper row) and syntaxin 1 (middle row), as well as melamine sections of cells expressing R-synuclein (lower row), with or without FP tags (see Materials and Methods for the procedures involved in generating membrane sheets). As for the previous figure, we would like to note that the Star635P fluorescence is shown and not the FP fluorescence. Scale bar, 500 nm. Bar graphs were generated for the median spot size and peak intensity ((SEM; black bars for untagged proteins, green bars for the FP-tagged ones). The graphs show averages of three independent experiments for SNAP25 and syntaxin 1, while four independent experiments contribute to the data shown for R-synuclein (two for the G141TAG mutant and two for the T142TAG). None of the differences were statistically significant (p > 0.05, Student's t test).

To test these results by a different imaging procedure, we analyzed three proteins that were often described to form multimolecular arrangements in stimulated emission depletion (STED) microscopy. For this purpose, we chose the transmembrane protein syntaxin 1, the membrane-attached protein SNAP-25, and the soluble protein R-synuclein. No significant differences were observed between their FP-tagged and nontagged variants in STED microscopy (Figure 5), in close agreement with the GSDIM results. Since STED is less restrictive on the imaging conditions, we could also use a different embedding procedure than in GSDIM. The membrane proteins (syntaxin 1, SNAP-25) were directly embedded in a fluid medium, Mowiol, rather than melamine (as in the case of GSDIM), and were afterward imaged without any further processing. This experiment therefore further verifies that a melamine-independent embedding procedure also results in indistinguishable FP-tagged and nontagged protein clusters. CONCLUSION We conclude that FP tagging did not generally result in major changes. For the few proteins that did change upon tagging, the direction is not unitary: four proteins formed larger and/or brighter assemblies, while the opposite took place for the remaining two. The lack of a generally observable effect is not due to the GSDIM (i.e., STORM-like) technique, since the study of three of the proteins by STED microscopy, at a comparable resolution, also revealed no significant differences between the FP-tagged chimeras and the untagged variants (Figure 5). Overall, our experiments suggest that, despite their relatively large size, the FP tags result in surprisingly limited changes in the clustering behavior of a variety of proteins. Therefore, the FP tagging does not automatically imply perturbation to the target protein's organization. Nevertheless, care must still be taken in expressing the target at biologically relevant levels, rather than overexpressing it. To enable experimenters to test their own proteins of interest, we include a detailed protocol for the comparison between FP-tagged and nontagged proteins, as part of the Supporting Information. Following this protocol, a decision can be made on whether a protein is affected by FP tagging within approximately 10 days. Finally, for those proteins whose function or organization is corrupted by FP tagging, we suggest that UAAs would be an efficient alternative. They are becoming widely available for nanoscopy and have the advantage of being more flexible than FPs in the choice of fluorophores and in the choice of the position VOL. XXX

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for difficult target proteins, at least for fixed-cell nanoscopy.

MATERIALS AND METHODS

directly to the sample stage in order to minimize sample drift. Sample focus was adjusted using a piezo-controlled objective positioner (Physik Instrumente). For GSDIM imaging, photoswitchable AlexaFluor647 was excited using 642 nm light and in some measurements was also exposed to 405 nm light to increase the activation rate of switching. A fiber laser (MPB Communications, 2RU-VFL-P-1000-642) was used to generate 642 nm light. The laser illumination was configured such that the illumination angle could be varied between an epi-illumination geometry and a total internal reflection (TIRF) illumination mode. For GSDIM data acquisition, the sample was illuminated with oblique illumination (not TIRF) to reduce background signal. Fluorescence emission of AlexaFluor647 was detected using an sCMOS camera (PCO Edge monochrome). GSDIM imaging was carried out according to the manufacturer's instructions. The camera exposure time was set to 10 ms, and the particle detection threshold was set to 25 in the Leica software (Leica Application Suite, Advanced Fluorescence, version 3.2). We used the “Auto Event Control” option of the Leica software, which increases the intensity of the 405 nm laser when the number of events recorded per image is too low. Therefore, the 405 nm laser power increased during the acquisition of the individual images from 0% to 100% of the maximum power. The 642 nm laser was used at 11% of the full power. Membrane Sheet Generation for STED Imaging. We used plasma membrane sheets, rather than melamine sections, for the STED imaging of two proteins that are found primarily on the plasma membrane, syntaxin 1 and SNAP-25. This procedure offers the same axial resolution as melamine sections in the case of plasma membrane proteins and is easier to perform. The cells were briefly washed with PBS, then sonicated in a 9 cm dish filled with ice-cold KGlu buffer (120 mM monopotassium glutamate, 20 mM potassium acetate, 2 mM EGTA, 20 mM HEPESKOH, pH 7.2). The sonication procedure was performed as follows: the coverslip was placed in the center of the dish at a distance of approximately 1 cm from the sonication tip. During sonication a 100 ms pulse was applied using a Branson Sonifier 450 (Branson Ultrasonics Corp., Danbury, CT, USA), equipped with a 2.5 mm sonication tip. Subsequently, the coverslip was briefly washed in ice-cold KGlu buffer and fixed. Click Reaction for STED Imaging. R-Synuclein samples were fixed, permeabilized, and embedded as whole cells as described for GSDIM preparations. The click labeling was performed in the presence of 50 μM Star635P-azide. Membrane sheets were fixed, quenched, and subjected to click reaction in a similar manner to whole cell samples but without the addition of Triton-X 100. In addition, the final click reaction mix contained 50 μM Star635P-azide instead of AlexaFluor647-azide. The sonicated samples were mounted in Mowiol (24% w/v glycerol, 0.1 M Tris-HCl, pH 8.5, 9.6% w/v Mowiol 4-88; Carl Roth GmbH, Karlsruhe, Germany). STED Microscopy. The images were taken with a STED microscope based on a fiber laser, emitting 1.2 ns pulses at a wavelength of 775 nm and 20 MHz repetition rate (IPG Photonics), to inhibit fluorescence by stimulated emission. See ref 51 for more details. Using a pulsed diode laser emitting 70 ps pulses at 640 nm wavelength, the fluorophores were excited in a diffraction-limited area. The 775 nm STED beam passes a 0
360 vortex linear phase ramp (RPC Photonics, Rochester, NY, USA) and a λ/4 plate to become circularly polarized, causing the focal beam intensity to resemble a “doughnut” having a central zero. The doughnut-shaped STED beam allows molecular fluorescence only in a subdiffraction-sized area around the zero intensity point. Pulse energies of 7.5 nJ for STED and 0.4 pJ for excitation (both measured at the back aperture of the objective lens) led to areas of approximately 30 nm in diameter, which also corresponds to the resolution obtained. Images were acquired by scanning the sample with a 3-axis piezo stage

A full description of materials, constructs, and cloning procedures is included in Supporting Information. Standard methods (epifluorescence microscopy, image analysis) are also presented in the Supporting Information. Transfection and UAA Incorporation. The cells were grown on poly-L-lysine-coated coverslips for 6
12 h to a 70
80% confluence. Approximately 1 h before transfection, the medium was supplemented with 250 μM propargyl-L-lysine (dissolved as a 1 M stock in DMSO). For incorporation to take place, the cells were transfected with vectors encoding for (i) the aminoacyltRNA-synthetase/tRNA pair (pCMV tRNA-Pyl RS WT)48 and (ii) the protein of interest (with or without a fluorescent protein tag), containing the Amber codon. The plasmids and the Lipofectamine 2000 reagent (Life Technologies) were separately equilibrated for 5 min in Opti-MEM (Gibco). Then the solutions were mixed and incubated for 20 min. The resulting mixture was applied to the cells, which were allowed to express the constructs for 18 h in a humidified incubator at 37 C. Two hours before sonication or fixation the medium was exchanged to normal Dulbecco's modified Eagle's medium (DMEM). Click Reaction for GSDIM Imaging. Whole cell samples were fixed for 30 min with 4% paraformaldehyde (PFA) in PBS (phosphatebuffered saline: 137 mM NaCl, 2.7 mM KCl, 10 mM Na2HPO4, 2 mM KH2PO4, pH 7.3) or with 0.2% glutaraldehyde-4% PFA in PBS. Then they were quenched for 20 min in 100 mM NH4Cl and 100 mM glycine (in PBS), followed by a brief wash with PBS. The cells were permeabilized with 0.1% Triton-X 100 in PBS, were incubated for 15 min with 5% BSA and 5% peptone in 0.1% Triton-X 100-PBS, and were briefly washed with 3% bovine serum albumin (BSA) in PBS before the click reaction. The click reaction mix contained 1 cell reaction buffer (component A), 2 mM CuSO4 (component B), a 10 dilution of Click-iT reaction buffer additive (component C), and a final concentration of 3 μM AlexaFluor647-azide in water. According to the manufacturer's protocol (Click-iT cell reaction buffer kit, Life Technologies), this click solution was freshly prepared and was added to the samples for 30 min in a dark humidified chamber at room temperature. The samples were afterward washed with PBS containing 5% BSA and 5% peptone (three solution exchanges, 5 min each), followed by three PBS washes (5 min each). The cells were then embedded in melamine. Plastic Embedding of Whole Cells and Thin Sectioning for GSDIM Imaging. The cells were embedded in melamine (2,4,6-tris[bis(methoxymethyl)amino]-1,3,5-triazine, TCI Europe, Zwijndrecht, Belgium) as previously described.49 The samples were first incubated with melamine for 24 h at room temperature (on silica gel, for dehydration purposes), to allow melamine penetration into the cells. They were then incubated for 24 h at 40 C and for a further 48 h at 60 C. The melamine-embedded samples where then cut into 100 nm sections using a microtome, and these were mounted on microscope coverslips. Imaging Buffer for GSDIM Imaging. All imaging experiments were carried out in MEA imaging buffer as previously described.50 The imaging buffer consists of 50 mM Tris-Cl (pH 8.0), 10 mM NaCl, 10% glucose (w/v), 10 mM β-mercaptoethylamine (pH 8.5, Sigma), and 1% of a stock solution of enzymatic oxygen scavenger system. The oxygen scavenging system was added to the buffer immediately before use. The oxygen scavenger stock solution was prepared by mixing glucose oxidase powder (10 mg, Sigma) with catalase (50 μL, 20 mg/mL, Sigma) in PBS (200 μL) and centrifuging the mixture at 13 000 rpm for 1 min. GSDIM Microscopy. All imaging measurements were performed using a commercial Leica GSDIM super-resolution microscope. The microscope was fitted with a 160 oil-immersion objective lens (Leica GSDIM), which enabled efficient detection of single fluorophores. This objective was mechanically coupled

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of the fluorophore within the protein sequence. This should allow UAAs to develop into the tools of choice

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Acknowledgment. The authors would like to acknowledge Dr. Vladimir Belov and Dr. Kirill Kolmakov (Department of NanoBiophotonics, Max Planck Institute for Biophysical Chemistry, Göttingen, Germany) for providing Star635P-azide and Prof. Dr. Reinhard Jahn, Prof. Dr. Thorsten Lang, Dr. John Chua, and Dr. Marcus Niebert for providing the constructs listed in the Supporting Information. We thank Christina Schaefer and Gaby Klaehn for technical support. I.C.V. has been supported by a Dorothea Schlözer fellowship. The work was supported by grants to S.O.R. from the European Research Council (ERC2013-CoG NeuroMolAnatomy) and from the Deutsche Forschungsgemeinschaft (DFG) SFB 889/A5. I.N. acknowledges financial support by the European Commission Seventh Framework Programme (FP7) through Marie Curie Actions (FP7PEOPLE-IEF) and a European Molecular Biology Organization (EMBO) long-term fellowship. E.A.L. acknowledges funding by the Emmy Noether program and SPP1623 of the DFG. S.O.R. and T.F.O. have been granted support from the Deutsche Forschungsgemeinschaft Cluster of Excellence Nanoscale Microscopy and Molecular Physiology of the Brain (CNMPB, EXC 171). Supporting Information Available: The Supporting Information is available free of charge on the ACS Publications website at DOI: 10.1021/acsnano.5b04434. Materials, cells, constructs, cloning strategy, epifluorescence methods, data analysis and statistics, Tables S1, S2, and S3, Figures S1 and S2, and the protocol for testing if other proteins are affected by FP tagging (PDF)

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and detecting fluorescence photons with an avalanche photo diode (Micro Photon Devices, Bolzano, Italy). A 690 ( 60 nm fluorescence filter prevented detection of photons emitted by the lasers. The pulse repetition rate of the excitation laser was set to 1.1 MHz, in order to reduce photobleaching by relaxation of the triplet state.52 Furthermore, an electrooptic modulator blocked the STED pulses that did not succeed pulses for excitation. The implementation of time gating passes only photons arriving within a time window of 1
10 ns after the excitation pulse to the computer, hence considerably reducing the dark count noise and avoiding the detection of spontaneously emitted photons during the action of the STED pulse. The pixel size and dwell time were set to 12 nm and 3 ms, respectively. Conflict of Interest: The authors declare no competing financial interest.

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