Here, we present a protocol for multi-color localization of single membrane proteins in organelles of live cells. To attach fluorophores, self-labeling proteins are used. Proteins, located in different membranes compartments of the same organelle, can be localized with a precision of ~18 nm.
Knowledge about the localization of proteins in cellular subcompartments is crucial to understand their specific function. Here, we present a super-resolution technique that allows for the determination of the microcompartments that are accessible for proteins by generating localization and tracking maps of these proteins. Moreover, by multi-color localization microscopy, the localization and tracking profiles of proteins in different subcompartments are obtained simultaneously. The technique is specific for live cells and is based on the repetitive imaging of single mobile membrane proteins. Proteins of interest are genetically fused with specific, so-called self-labeling tags. These tags are enzymes that react with a substrate in a covalent manner. Conjugated to these substrates are fluorescent dyes. Reaction of the enzyme-tagged proteins with the fluorescence labeled substrates results in labeled proteins. Here, Tetramethylrhodamine (TMR) and Silicon Rhodamine (SiR) are used as fluorescent dyes attached to the substrates of the enzymes. By using substrate concentrations in the pM to nM range, sub-stoichiometric labeling is achieved that results in distinct signals. These signals are localized with ~15–27 nm precision. The technique allows for multi-color imaging of single molecules, whereby the number of colors is limited by the available membrane-permeable dyes and the repertoire of self-labeling enzymes. We show the feasibility of the technique by determining the localization of the quality control enzyme (Pten)-induced kinase 1 (PINK1) in different mitochondrial compartments during its processing in relation to other membrane proteins. The test for true physical interactions between differently labeled single proteins by single molecule FRET or co-tracking is restricted, though, because the low labeling degrees decrease the probability for having two adjacent proteins labeled at the same time. While the technique is strong for imaging proteins in membrane compartments, in most cases it is not appropriate to determine the localization of highly mobile soluble proteins.
The goal of this protocol is to provide an imaging method to localize and track single membrane proteins inside live cells. We call this method Tracking and Localization Microscopy (TALM)1,2. Like Stochastic Optical Reconstruction Microscopy (STORM)3 and Fluorescence Photoactivation Localization Microscopy ((F)PALM)4,5, TALM is a single molecule-based fluorescence localization technique. However, it is distinct in the way that the mobility of membrane proteins in combination with repetitive imaging of the same labeled molecule at different positions reveals the microcompartment that is accessible for the mobile protein. In other words, the possible localizations of the protein are set by the architecture of the organelle and by the mobility of the protein1. The method is complementary to various other super-resolution techniques6,7,8 because it reveals localization and trajectory maps by imaging mobile proteins. The labeling is based on using genetically engineered fusion proteins that are per se non-fluorescent. These fusion proteins are self-labeling enzymes that react covalently with a substrate conjugated to a dye. This procedure has the advantage that the labeling degree can be controlled by the amount of added substrate. Furthermore, it allows to vary the color of fluorescence, depending on the chosen conjugated dye. Several self-labeling enzyme-tags are available9. Another advantage of using self-labeling enzyme-tags is, that the conjugated dyes usually are more stable and brighter than fluorescent proteins1 and individual proteins therefore can be recorded longer and more precisely until they are bleached. This allows for the recording of trajectories of mobile proteins and the extraction of diffusion coefficients10,11.
Here, we demonstrate the feasibility of TALM with mitochondrial membrane proteins, but it can also be applied for other intra- and extra-cellular membrane proteins, including different cell types12,13. We show that multi-color TALM further allows for the simultaneous distinction of proteins in different subcompartments in complementation to existing super-resolution fluorescence microscopy techniques14,15,16. TALM is compatible with live cell imaging17. The photo-physics of the chosen rhodamines Tetramethylrhodamine (TMR) and Silicon-Rhodamien (SiR), in particular their brightness and stability, allows to record single membrane proteins over multiple frames providing localization (and trajectory) maps. However, TALM is limited for the localization of soluble proteins with high diffusion coefficients since the motion blur is too high and the collected photons per frame are too low for proper localization. Besides, TALM requires less excitation power than for example STORM or Stimulated Emission Depletion (STED) microscopy6,7, reducing phototoxic effects. This is important, since phototoxic stress often affects organellar morphology18 and thus mobility analysis19. In sum, we present multi-color TALM in living cells as a technique that fills a gap between the localization microscopy methods STORM/STED/(F)PALM and techniques that analyze protein mobility such as fluorescence recovery after photobleaching (FRAP)20,21, fluorescence correlation spectroscopy (FCS)22, and fluorescence cross correlation spectroscopy (FCCS)11,23.
The following protocol follows the guidelines of the local institution research ethics committee.
1. Methods
2. Microscopy
Figure 1: Optical layout for multi-color tracking and localization microscopy (TALM) with orange and red emitters. (A) Inverted microscope setup with at least two excitation lasers, a TIRF condenser, a TIRF suitable objective, an image splitter, and a sensitive camera. Inset: to excite organelles inside cells, the angle of the incident beam must be set smaller than the critical angle for TIRF to achieve highly inclined and laminated optical sheet illumination (HILO). DC1: Dichroic mirror 1; DC2: Dichroic mirror 2. EF: emission filter. (B) Test on optical drift by imaging positions of a fluorescent bead for 10,000 frames with the same frame rate as of the following experiments (here: 15 Hz). Connected positions of the first 500 frames and the last 500 frames show the drift. Also, a merged image with the position of the first and the last frame in red and blue show a minimal drift. The drift is the distance between the centre of the signals divided by the total recording time, here 125 pm/s. (C) Check on the clear separation of signals, here TMR and SiR. For both channels, cumulative sum images from 3,000 frames (TMR in Channel 1 and SiR in Channel 2) were generated. SiRHTL was attached to Tom20-HaloTag and TMRHTL to OxPhos complex V-HaloTag. Colors are false colors. Scale bars = 100 nm (B) and 1 µm (C). Please click here to view a larger version of this figure.
Figure 2: Workflow for dual color alignment. (A) The coverslip with the fluorescent beads is mounted in a sample holder between a PTFE and a rubber ring. Then the upper and lower part of the chamber are bolted together. (B) Physical alignment of the channel views that are generated by the image splitter. Recorded fluorescent signals from beads (0.1 µm) in two channels (green and red, false colors) are merged. The corresponding screws at the optical image splitter are manually turned until the best overlay of the different signals is achieved (yellow color, lower panel). (C) Generation of a transformation matrix for post-processive channel alignment. For precise localization of a particle, it is necessary to determine the point spread function (PSF) in dependence on the emission wavelength and the numerical aperture of the objective. The center of a PSF can be determined by its intensity profile analyzed by a symmetric two-dimensional Gaussian fit. The resulting localization of the signal peak is then projected on the original, blurred signals. In a merged image, the localized centers of the signals from the two channels are connected to generate a transformation matrix that is later used for the post-processive alignment of the experimental data. Scale bars = 1 µm (B, C). Please click here to view a larger version of this figure.
Figure 3: Steps during single molecule localization microscopy. (A) A coverslip with the specimen is mounted between the top and bottom part (grey) of the homemade sample holder (designed by J. Bereiter-Hahn). A rubber ring (red) and a PTFE ring (white) seal the system from above and below the coverslip, when the sample-holder parts are bolt together. (B) Signal to noise ratio of the TMR signal. (C) Calculated localization precision histogram for all localized particles. (D) Choice of a reasonable region for imaging, here, the cell periphery with clearly separated mitochondria. (E) Recording and image processing: a single frame with distinct single molecule signals is shown (here, single molecules of CV-HaloTag/TMRHTL were recorded). (F) Intensity of TMR over the recording time. (G) Cumulative sum image of 3,000 frames, unprocessed. (H) Particles of CV-HaloTag/TMRHTL localized with a 2D Gaussian function from a single frame. (I) Cumulative, rendered sum image showing all localized CV-HaloTag/TMRHTL particles from 3,000 frames. Please click here to view a larger version of this figure.
Multi-color imaging and colocalization analysis can help to determine the sub-organellar localization of proteins. We demonstrated this earlier with the cytosolic phosphatase and tensin homologue, PINK1, that has different sub-mitochondrial locations due to its processing by mitochondrial proteases17. PINK1 is an important factor guaranteeing mitochondrial functionality34,35. To determine the localization of PINK1 in different mitochondrial compartments in the course of its processing (Figure 4A), multi-color super-resolution microscopy with live cells was performed. Therefore, PINK1 was genetically fused to a self-labeling tag (HaloTag). To determine its localization relative to other proteins in functional and dysfunctional mitochondria, Tom20, as part of the translocase of the outer mitochondrial membrane (TOM)36, was tagged with another self-labeling tag (SNAP-tag). Time series with at least 1,000 frames (96 frames per s) were recorded. The cumulative image of all signals over time from one channel showed that PINK1 was localized in the cytosol and in the mitochondria (Figure 4B, left panel) under normal conditions, while it was retained at the outer membrane of depolarized mitochondria (treatment with 10 µM of the uncoupler carbonyl cyanide m-chlorophenyl hydrazine, CCCP, for 20 min). The corresponding trajectory maps also show this difference in the spatio-temporal organization. As the summed images of localized particles reveal, PINK1 is distributed mainly inside polarized mitochondria (Figure 4C, left upper panel), while Tom20 is localized in the outer mitochondrial membrane (OM) (Figure 4C, left lower panel). This becomes even more clear in the merged image of localized Tom20 and PINK1 particles, where the distribution of the outer membrane protein Tom20 is much broader (Figure 4C, right panel).
Figure 4: Dual color super-resolution microscopy showing the subcompartmental localization of PINK1 in mitochondria. (A) Hypothesized PINK1 shuttling and processing in mitochondria. (B) Localization and trajectory maps of PINK1 under normal conditions and in depolarized mitochondria. (C) Dual-color super-resolution localization microscopy to reveal mitochondrial localization of PINK1 in relation to Tom20. Image series of 1,000 frames recorded with a frame rate of 96 frames per s. Left upper panel: Localization map of PINK1 (labeled via HaloTag with TMRHTL). Left lower panel: localization map of Tom20 (labeled via SNAP-Tag with SiRBG). Right panel: Merge cumulative sum image of PINK1-TMR (red) and Tom20-SiR(blue)localizations. (D) Same data plus localizations of respiratory complex I (CI, green) in the inner mitochondrial membrane. CI was fused to paGFP (CI-paGFP). Right: Mean cross section profile of Tom20 (blue), PINK1 (red), and CI (green) fluorescence in the marked region of the merged image. The mean cross section profiles were obtained by averaging up to 30 parallel oriented cross lines (interval 40 nm). Adapted version reprinted with permission from Beinlich et al.16, copyright (2015) ACS chemical biology. All colors are false colors. Scale bars = 1 µm. Please click here to view a larger version of this figure.
To verify the localization of PINK1 inside mitochondria, a third protein localized in the inner mitochondrial membrane was also imaged. Therefore, the 30 kDa subunit of respiratory complex I (CI) was fused to photoactivable GFP (paGFP) and recorded by FPALM4. The cumulative triple color image and the cross-section distribution show the overlay of CI (green) and PINK1 (red) (Figure 4D), confirming the import of full-length PINK1. PINK1 has an N-terminal mitochondrial targeting sequence and thus the label was put at the C-terminus of the kinase. Co-localization with CI demonstrates that the full length PINK1 had been imported. The fluorescence distribution along a cross-section shows that a quota of PINK1 particles apparently also colocalized with Tom20 in the outer mitochondrial membrane. Probably, this is PINK1 associated with Tom20, which is the import receptor. As these data demonstrate, several mitochondrial micro-compartments are occupied by subpopulations of PINK1. To address this question of sub-mitochondrial localization by biochemical methods would be much more difficult, since it would require stringent sub-fractionation of the different membranes and a clear separation of soluble components to exclude cross-contamination.
Supplementary Video 1: Footage of single molecule recording, 100 frames, CV-HaloTag/TMRHTL. Please click here to download this file.
Supplementary Video 2: Localized data. Please click here to download this file.
Here, a technique for dual color single molecule localization of mobile membrane proteins was presented. Following the protocol, membrane proteins are fused to self-labeling proteins that react with the rhodamine dyes TMR and SiR conjugated to their respective substrates. Rhodamine dyes are bright and photostable and thus allow for repetitive imaging1. For successful performance, several conditions and critical topics have to be kept in mind.
First, it is important to choose appropriate filters and splitters to separate the signals from TMR and SiR clearly. To reduce background from outside the cell, a precleaning and coating of the coverslip with PLL-PEG-RGD was helpful. The final concentration of the dye-substrate must be tested by considering the affinity of the self-labeling enzyme for its substrate and the accessibility of the enzyme inside the cell. Nano- to picomolar concentrations were sufficient for mitochondrial proteins17 and stress granules13. For other organelles, the required concentrations need to be tested. If the staining is too strong at the beginning of the recording and no single molecules are distinguishable, it is best to wait until bleaching has reduced the amount of fluorescent dyes so that single particles (SP) can be discerned. We have found that concentrations of dyes higher than 30 nM for BG-substrates and 1 nM for HTL substrates during the staining procedure, or longer staining times in order to increase the number of labeled molecules, are not feasible. For each experiment, the intensity of the excitation laser has to be adapted, since the subcellular location of the dye influences its fluorescence behavior (quantum yield, bleaching, blinking)5 due to different environmental conditions such as pH and redox state3. It further must be mentioned that the background is higher in dual-color experiments when compared to single color imaging. This influences the precision of localization. Therefore, it is critical to optimize the laser power to achieve good signal to noise (S/N) ratios but low photobleaching rates. Typical laser power densities in HILO microscopy are in the range of 25 ± 8 kW/cm2. For fine-tuning of excitation power, different sets of neutral density filters can be used if laser diodes are not directly modulated and not modulated via an acousto-optical modulator. It is recommended to use special 2 mm thick mirrors for TIRF to reduce beam distortions. Especially under TIR and HILO conditions, very efficient blocking of excitation light is needed. For single molecule imaging and localization with subpixel accuracy, a proper spatial sampling frequency (Nyquist-Shannon sampling theorem) is required. This should be twice as high as the maximum spatial frequency defined by the resolution limit of the imaging system37. Using an oil immersion objective with a high numerical aperture (NA >1.4), the resolution, d = λ/(2 × NA), is typically at 200-250 nm for orange to far-red dyes. Thus, considering the physical pixel size of the detector, the magnification of the imaging optics should result in an image pixel size of approximately 100 nm. For an EMCCD camera with a pixel size of 16 x 16 µm2 in combination with a 150X objective, the pixel size is 106.7 nm and thus adequate.
Generally, it is suggested to use stable transfected cells because this has the advantage that a steady amount of tagged proteins is expressed in most of the cells24. However, for dual-color localization and tracking experiments, this would require a double transfection and double selection with different antibiotics. Therefore, it is often more feasible to transiently transfect an already stable cell line with a second plasmid encoding the additional tagged protein of interest.
For mitochondria, movement and fragmentation is a critical issue. Fragmented or moving organelles should not be recorded or analyzed. Movement can be checked by overlaying rendered images of localized molecules from the beginning and the end of the experiment, with two different (false) colors. A homogeneous distribution of the signals from the organelles in the overlay indicates that organelles have not moved. Generally, room temperature reduces the movement of cellular compounds and structures.
The authors have nothing to disclose.
The authors would like to thank the Biophysics group and Jacob Piehler at the University of Osnabrück for continuous support, Wladislaw Kohl for technical assistance and preparation of material, and the CellNanOs board for providing microscopes for use. The project was funded by the SFB 944.
(2-(4-(2-hydroxyethyl)-1-piperazinyl)-ethanesulfonic acid, 1 M) (HEPES) | Biochrom | #1104E | |
DC1: Dichroid beam splitter | Chroma | 640 dcxr | NC506031 |
DC2: Polychroic Mirror, beamsplitter | Chroma | zt405/488/561/640rpc | discontinued |
Dulbecco´s Phosphate-Buffered Saline (PBS) 1x (w/o Ca & Mg) | Sigma-Aldrich & Co. | #RNBF8311 | |
Earle´s MEM without phenol red, without L-Glutamine and without NaHCO3 containing 1% FBS, 0.1% HEPES, 0.1% NEAA, 0.1% Alanyl-L-Glutamine and 34.78% sodium hydrogen carbonate (NaHCO3 0.75g/l) | Imaging medium | ||
Earle´s minimum essential medium (MEM) with phenol red, containing 1% Fetal Bovine Serum Superior (FBS), 0.1% HEPES (2-(4-(2-hydroxyethyl)-1-piperazinyl)-ethanesulfonic acid, 1 M), and 0.1% non-essential amino acids (NEAA) | Growth medium | ||
EF: Emission filter quadbandpass | AHF analysentechnik | F72-866 | Brightline HC 446 nm/523 nm/600 nm/677 nm |
EMCCD camera | Andor | Andor iXON 897 | EMCCD camera |
Emission filter QuadView filter cubes, orange | AHF analysentechnik | F39-637 | bandpass 582 – 619 nm |
Emission filter QuadView filter cubes, red | Chroma | bandpass 655 – 725 nm (HQ 690/70) | |
FBS (Fetal bovine serum) superior | Biochrom | S0615 | |
Fluorescent beads: TetraSpeck™ Microspheres, 0.1 µm, fluorescent blue/green/orange/dark red | Thermo Fisher Scientific | T7279 | fluorescent microspheres |
Glutamine | Biochrom | #0951C | |
HeLa cells | DSMZ | ACC-57 | Cervical carcinoma cells from patient Henrietta Lacks |
Hela cells CI::paGFP, stable | Muster et al., PLOSOne 2010 | ||
Hela cells CV g::Halo7-Tag, stable | Appelhans et al., NanoLett 2012 | ||
Hela cells Tom20::Halo7-Tag, stable | Appelhans et al., NanoLett 2012 | ||
Image splitter | Photometrics | Dual-View QV2 | image splitter emission |
Imaging processing software | ImageJ2 / Fiji | freeware | |
Immersion Oil – ImmersolTM 518 F (ne = 1.518, ve = 45) | Carl Zeiss Jena GmbH | 444960-0000-000 | |
Inverted epifluorescence microscope | Olympus IX-71/73/83 | ||
Laser 561 nm, 200 mW | CrystaLaser | CL-561-200 | 561 nm emission |
Laser 642 nm, 140 mW | Omicron | Luxx-642-140 | 642 nm emission |
MATLAB | MathWorks | version R2013a | |
MEM with Earle's Balanced Salt Solution 2.2 g/L NaHCO3, stable glutamine w/o PR | Biochrom | FG-0385 | |
MEM with Earle's Balanced Salt Solution with 2.2 g/L NaHCO3, stable glutamine, Phenolred | Biochrom | FG-0325 | |
MitoTracker® Deep Red FM | Thermo Fisher Scientific | M22426 | dye |
MitoTracker® Green FM | Thermo Fisher Scientific | M7514 | dye |
Multi-mode-optical polarization maintaining monomode fiber | Pointsource/Qioptiq | KineFLEX | |
NHS-PEG-MAL, Rapp Polymer | Rapp Polymere GmbH Tübingen | coverslip coating | |
non-essential amino acids (NEAA) | Biochrom | #0802E | |
PEG 800 (Polyethylene glycol) 10 % | Carl Roth GmbH | Art No. 0263.1 | coverslip coating |
Penicillin/Streptomycin | Biochrom | #0122E | |
Plasmid for PINK1-Halo7-Tag expression | Beinlich et al., ACS Chemical Biology 2015 | ||
Poly-L-lysine (1.2 mg/ml) | Sigma-Aldrich & Co. | Cat. No.P9155 | coverslip coating |
RGD Peptide (Ac-CGRGDS-COOH) | Coring System Diagnostix GmbH, Gernsheim | coverslip coating / Intergrin receptor motif | |
Silicon Rhodamine linked to HaloTag®-Ligand (SiRHTL) | personal gift from Kai Johnson | dye | |
Software analysis plugin | self-written C. P. Richter, Biophysik Osnabrück | SLIMFAST 16g | |
Tetramethylrhodamine / SNAP-Cell® TMR-Star linked to SNAP-Ligand (TMRstar) | New England Biolab® | S9105S | dye |
Tetramethylrhodamine linked to HaloTag®-Ligand (TMRHTL) | Promega | G8251 | dye |
TIRF condensor | Olympus | Cell^TIRF MITICO System | TIRF condensor |
TIRF microscope controlling software | Olympus cellSens 1.12 | ||
TIRF objective | Olympus | 150x oil objective (N.A. 1.45; Olympus UAPO) | |
Trypsin/EDTA 10x | Biochrom | #0266 | |
Water H2O 99,5 % Rotipuran® Low organic | Carl Roth GmbH | Art. No. HN57.1 |