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The Use of miniSOG in the Localization of Mitochondrial Proteins
CHAPTER TEN
The Use of miniSOG in the Localization of Mitochondrial Proteins Guy A. Perkins1 National Center for Microscopy and Imaging Research, University of California, San Diego, California, USA 1 Corresponding author: e-mail address: [email protected]
Contents 1. 2. 3. 4. 5.
Introduction Requirements for CLEM Labeling miniSOG Features Resolution Photooxidation Protocol for a Monolayer of Cultured Cells 5.1 Chimera construction and transfection notes 6. Photooxidation Protocol for Tissues 7. Example of miniSOG Use with MCU 8. Conclusions and Future Work Acknowledgments References
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Abstract In this chapter, we provide details along with considerations and future directions for the use of miniSOG (for mini Singlet Oxygen Generator), a versatile label for correlated light and electron microscopy of genetically tagged proteins in cells, tissues, and organisms. This new visualizable genetic tag improves the ability of biologists to locate specific proteins at nanoscale resolution and to see these tagged proteins in the environment of structural landmarks that we are used to navigating by, such as mitochondrial membranes and compartments. miniSOG provides high-quality ultrastructural preservation and permits three-dimensional protein localization via electron tomography or serial section block-face scanning electron microscopy. miniSOG is now doing for electron microscopy what the family of green fluorescent protein did for fluorescence microscopy.
Methods in Enzymology, Volume 547 ISSN 0076-6879 http://dx.doi.org/10.1016/B978-0-12-801415-8.00010-2
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2014 Elsevier Inc. All rights reserved.
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1. INTRODUCTION CLEM—correlated light and electron microscopy (the same area examined by both microscopy techniques) or correlative light and electron microscopy (the same specimen, but not the same area)—is an expanding tool for dissecting the protein landscape useful for providing clues to cell and tissue function at high spatial resolution (Ellisman, Deerinck, Shu, & Sosinsky, 2012). Each imaging modality provides unique information, and the combination of light and electron microscopies can contribute to a better understanding of the spatiotemporal patterns of protein expression, trafficking, binding partners, and function. Fluorescence light microscopy provides the dynamics and options for multiprotein labeling, and electron microscopy provides the increased resolution and cellular context. The use of genetically appended tags to specific proteins to be able to localize them in their complex cellular milieu is growing as new tags have been developed.
2. REQUIREMENTS FOR CLEM LABELING Certain requirements need to be met for a genetic tag to be useful as a CLEM label. First and foremost, the genetic sequence of the protein of interest must be known. Second, a genetic tag must be nontoxic to the cell in which it is expressed. It should be active in the physiological environment where expressed. In addition to the exhibiting fluorescence, it should either independently or with additional ligands or cofactors create an electrondense label for CLEM imaging. A genetic tag has the additional attractive property that it is stoichiometric to its target protein. This quality can be important for quantification of low copy number proteins within cells. As with any exogenously expressed protein, it is important to conduct control experiments to ensure that recombinant proteins are expressed and localized the same as endogenously expressed proteins. A commonly used control is comparison to immunolabeled native systems.
3. miniSOG FEATURES Mini Singlet Oxygen Generator (miniSOG) is one such recently engineered genetic tag. It is a 106-amino acid green fluorescent flavoprotein generated from Arabidopsis phototropin2 that efficiently generates singlet oxygen when illuminated with blue light (Shu et al., 2011). To generate
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Figure 10.1 When excited by blue light (dark gray in print version), miniSOG produces singlet oxygen, a reactive and short-lived excited state of O2. This singlet oxygen quickly oxidizes and polymerizes diaminobenzidine (DAB) into a localized osmiophilic density. Addition of osmium tetroxide to the polymerized DAB produces an electron-opaque stain that then allows nanometer-scale imaging of the tagged protein by electron microscopy.
singlet oxygen, miniSOG requires a flavin mononucleotide (FMN) cofactor. FMN, derived from the vitamin riboflavin, is endogenous to cells and is ubiquitous. It is indispensable in mitochondrial electron transport, fatty acid oxidation, and vitamin metabolism. Singlet oxygen, a reactive and short-lived excited state of O2, quickly oxidizes and polymerizes diaminobenzidine (DAB) into a localized osmiophilic mass, which osmicated density is clearly seen in the electron microscope (Fig. 10.1). Diffusion of this dense reaction product is minimized by extensive chemical cross-linking by glutaraldehyde prior to the generation of the reaction product. Because miniSOG is relatively small, about half the size of GFP, the protein tagging is less likely to influence protein targeting to the right location than larger genetic tags. Because miniSOG is genetically fused to the protein of interest and all other components (DAB, light, O2, and OsO4) are small and readily permeate well-fixed tissue, there is no conflict between labeling efficiency and established methods for excellent structural preservation. Moreover, miniSOG can photooxidize DAB within relatively thick slices of tissue, expanding its applicability. Indeed, miniSOG fusions have been expressed in adult mice using in utero electroporation. miniSOG causes no obvious cellular toxicity in the absence of light. Table 10.1 summarizes the properties of the miniSOG genetic label. The original report of a high quantum yield of singlet oxygen (0.47) has recently been challenged in two reports (Pimenta, Jensen, Breitenbach, Etzerodt, & Ogilby, 2013; Ruiz-Gonzalez, Cortajarena, Mejias, Agut, Nonell and Flors,
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Table 10.1 miniSOG Properties 1
Size (kDa)
Excitation (nm)
Emission (nm)
O2 Quantum yield
Fluorescent quantum yield
Brightness (M21 cm21)
13.9
447/473 (shoulder)
500/528 (shoulder)
0.47 (0.03a)
0.37
5820
a Pimenta et al. (2013) and Ruiz-Gonzalez et al. (2013) report a lower quantum yield of 0.03. Adapted from Shu et al. (2011) and table 1 of Wingen et al. (2014). Additional spectroscopic information is provided by List et al. (2014).
2013) that measured a quantum yield of 0.03 for miniSOG. The difference was attributed to the finding that miniSOG oxidizes ADPA, the standard singlet oxygen sensor used by both singlet oxygen-dependent and -independent processes. Interestingly, the low intrinsic quantum yield is compensated by a photoinduced transformation of miniSOG by cumulative irradiation that increases its quantum yield about 10-fold, which corroborates its utility for CLEM. miniSOG fusions have been made for many well characterized proteins and have been found to localize correctly in cultured cells, tissues, and intact organisms enabling CLEM without the need for exogenous ligands, probes, or destructive permeabilizing detergents. Therefore, miniSOG can be used for high-resolution 3D electron microscopy modalities, such as electron tomography and serial block-face scanning electron microscopy. This combination of high-resolution protein labeling and 3D electron microscopy now permits scientists to pursue answers to questions previously impossible to ask, such as does a particular mitochondrial protein localizes to the crista junction? Unlike particulate markers such as protein-conjugated immunogold or quantum dot labels, miniSOG does not just decorate the targeted protein but also is similar to a negative stain in that it outlines the protein (Ludwig et al., 2013). miniSOG can be used alone or in combination with different fluorescent proteins as the optical reporter for the tagged protein. Because miniSOG photobleaches rather quickly it thus may be necessary to fuse it to a second, nonoverlapping fluorescent genetic marker such as mCherry, tdTomato, or mKO2.
4. RESOLUTION The ultimate resolution achievable by miniSOG labeling remains to be determined. However, the diffusion of the reacted DAB product away from the miniSOG tag has been has been shown to be minimal, thus
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allowing proteins to be localized within a few nanometer precision (Ludwig et al., 2013). Using line scans across images of miniSOG-labeled caveolar coat protein, this group found that the polymerized and osmicated DAB electron density clearly resolved periodic density changes of about 10 nm. To accurately determine the resolution, one could express miniSOG on a subunit of a periodic solved structure, such as an isoform of actin or tubulin or an actin-associated protein. High-resolution electron tomography would then be performed. Subvolume averaging of tomographic reconstructions would define the molecular envelope of the tagged protein, which would then allow for the determination of the location of the osmium DAB density relative to the protein.
5. PHOTOOXIDATION PROTOCOL FOR A MONOLAYER OF CULTURED CELLS 5.1. Chimera construction and transfection notes Fusion constructions and transfections that work for GFP-family labeled proteins usually work for miniSOG fusions. Chemical transformation or lentiviral infections are often the most convenient, including stereotaxic injection of a lentivirus into a specific organ. However, electroporation is the method of choice for large plasmids. miniSOG fusions can be cloned into the pcDNA3.1 vector (Invitrogen). miniSOG or chimera cDNAs can be transfected into the cell line of choice using the Fugene (Promega) kit. Successful transfection of neurons has been achieved using Amaxa electroporation (Lonza AG, Germany). For rodents, miniSOG fusion constructs can be delivered into the lateral ventricle of embryos by in utero electroporation. It is recommended that two people be involved in the experiment to share the making of solutions, and performing the incubations and washes in order to not leave the samples in the various solutions too long. When removing solutions, it is important not to touch the central circle of the dish where the glass coverslip is as this will be the region imaged with CLEM. When adding solutions, it is important not to direct the stream of liquid on this central portion of the dish, but rather to the side of the dish so as not to disturb the cells. Note that many of the chemicals used are toxic, and so appropriate lab safety procedures should be followed. 1. Fix transfected cells cultured on glass bottom culture dishes (P35G-014-C, MatTek Corp., Ashland, MA) with 2% glutaraldehyde + 2 mM CaCl22H2O in 0.1 M sodium cacodylate buffer (pH 7.4) for 60 min.
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Warm fixative to 37 C. After removing the growth medium from these samples by either pouring off or pipetting off around the dish edge with a plastic disposable pipet, add the warmed fixative. Note that a glass pipet should not be used because microshards of glass may flake off and if embedded in the plastic resin may damage a diamond knife used for sectioning. Let the samples cool to room temperature for 5 min. Place the samples on ice for the last 55 min. Note that all aqueous solutions should be made with double-distilled water (ddH2O). From this step forward, it is important to keep the samples cold at all times until the 100% ethanol step below. Make up the cacodylate washing buffer and blocking solution (next 2 steps). Get them cooling on ice. It is wise to look one to two steps ahead to have chilled solutions ready when needed. 2. Wash samples with ice-cold 0.1 M sodium cacodylate buffer (pH 7.4) five times for 2 min each (5 2 min). Keep samples on ice. It is important that each sample not dry out by being uncovered by solution for more than a few seconds. So, when washing or replacing solutions, add the new solution quickly after removing the previous and do this for each sample before going to the next sample, i.e., do not remove the solution from all the samples before adding the new solution to all the samples. 3. Photoconversion block: Block samples for 15–20 min with ice-cold blocking solution consisting of 20 mM glycine, 10 mM potassium cyanide, and 10 mM aminotriazole (catalase inhibitor) in 0.1 M sodium cacodylate buffer. Right before use, add hydrogen peroxide: 1 μL per 25 mL of blocking solution. Keep samples on ice. 4. Wash samples with ice-cold 0.1 M cacodylate buffer 5 2 min. Keep samples on ice. Optional: Additional blocking step with mersalyl acid to reduce background nonspecific labeling of mitochondria. Note that the mitochondrial electron transport chain can generate singlet oxygen. Mersalyl acid is a mercury compound that poisons the ETC to block this singlet oxygen production. a. Block samples in 5–10 mM mersalyl acid (Sigma Cat. No. M9784-1G) for 30 min on ice. b. To make 10 mL of a 5 mM solution Dissolve 24 mg (50 μM) of the mersalyl acid in 50 μL of a 1 M NaOH stock solution. Vortex until almost all the mersalyl acid has been dissolved. Add 6.7 mL ddH2O + 3.3 mL of 0.3 M
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cacodylate buffer. Add 35 μL of a 1 M HCl stock solution. Filter with a 0.22 μm Millipore filter attached to a syringe. Cool this solution on ice before adding to the samples. 5. Wash samples in ice-cold 0.1 M cacodylate buffer 5 2 min. Keep samples on ice. 6. Make the DAB solution: DAB (free base—MW 214.27) (Cat. No. D8001-10G Sigma) Dissolve 0.005 mg free base DAB in 1 mL 0.1 M HCl (final concentration 25 mM stock solution) by vortexing until dissolved (may take a couple of minutes). Add 9 mL of 0.1 M cacodylate buffer. Filter solution into a new 20 mL scintillation vial (Fisher Scientific) with a 0.22 μm Millipore filter attached to a syringe. Photooxidation steps 1. Use an inverted confocal or inverted fluorescence microscope to identify transfected cells and for correlative light microscopic imaging. An inverted microscope ensures direct open access to the DAB solution. Set the microscope stage temperature to 4 C. 2. Take initial picture with minimal exposure to avoid photobleaching of the target area on the sample dish using both fluorescence and DIC. 3. Remove buffer from the dish and add enough DAB solution to cover the bottom of the dish. Let sit for 5 min. 4. Take off the dish lid. Gently blow oxygen continuously from an oxygen tank onto the sample over the top of the liquid surface. Alternately, the DAB solution on ice can be bubbled with oxygen and the solution in the dish refreshed every 1–2 min. 5. Illuminate the target area using a standard FITC filter set (EX470/40, DM510, BA520) with light from a 150 W xenon lamp to photooxidize until the sample turns a light brown as monitored by transmitted light. It is important to know that photooxidation is fast (typically 2–6 min) depending on how concentrated the labeled protein is and how accessible the miniSOG is in the folded protein. It is recommended that optimal timing to produce a well-visualized reaction product, but not to overreact, be determined by screening a small series of time points. If the illumination was too long, background density is observed in the electron microscope. Care should be taken not to illuminate with too intense beam as this may cause photodamage to cells. A setting of 30% power is typically used for the xenon lamp. Mitochondria are particularly sensitive and the damage most often seen is swollen matrix or matrix with regions devoid of cristae.
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6. Take final fluorescence and DIC pictures after photooxidation. Go to the next area in the same dish and repeat, usually three to four areas per dish. 7. Remove the sample from the microscope and wash it in ice-cold 0.1 M cacodylate buffer 5 2 min. Keep samples in buffer on ice until done with all plates. 8. Incubate samples in 1% osmium tetroxide (Electron Microscopy Sciences) in 0.1 M cacodylate buffer 30 min on ice. Work in the hood because osmium tetroxide is toxic and volatile. Electron microscope sample preparation steps 1. Wash samples in ice-cold ddH2O 3 2 min. Keep samples on ice. 2. Stain en bloc with 2% aqueous uranyl acetate (Ted Pella Inc.) 1 h on ice. If needed, this step can go overnight. Simply place the dishes in a refrigerator. Note that the first time doing the experiment it is recommended to have one plate sans uranyl acetate as it may mask the true signal, which is a higher concentration (contrast) of osmium bound to the DAB, especially relevant if the protein is sparse. 3. Dehydrate the samples in an ice-cold graded ethanol series (20%, 50%, 70%, and 90%) 2 min each. 4. Further dehydrate in 100% anhydrous ethanol 3 2 min at room temperature. Remove as much of the liquid as possible from each dehydration by pipet. Make sure that no water gets inside the dish during these dehydrations, and, of course in subsequent steps. Keep the lid on the dish to minimize water vapor on the samples. Try not to breathe over the dishes to minimize water vapor. Remove the ice bucket from the vicinity of the dishes to avoid water condensation close to the samples. From here to the end of the procedure, water is the enemy. 5. Infiltrate the samples in Durcupan ACM resin (Electron Microscopy Sciences) using 1:1 anhydrous ethanol:resin for 30 min at room temperature. The ethanol acts as a vehicle to aid the viscous resin to infiltrate into the cells. The four different bottles used are labeled A (or A/M), B, C, and D, respectively. The ratios of the Durcupan components should be 11.4 mL A:10 mL B:0.3 mL C:0.1 mL D. Because certain components are viscous, measure 11.4 g of A and 10 g B on a scale. For C and D, use 1 mL syringes to draw the solution from the bottle. Component C is smelly, so work in the hood. 6. Infiltrate the samples in 100% Durcupan ACM resin 3 1 h with freshly made resin each time because the resin hardens with time and this hardening impedes infiltration. Remove as much of the previous resin as
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possible taking care not to touch the central circle of the dish, where the glass cover slip is glued. At this point, ethanol is the enemy as even a little residual ethanol in the resin will make the samples soft and thus hard to section. 7. Polymerize the samples in an oven set to 60–70 C for 48 h. After the samples have cooled to room temperature, they should be “rock-hard,” important for good sectioning. Electron microscopy 1. Remove the glass coverslip from the bottom of the dish. It is usually difficult to remove this coverslip, and thus the strategy is to employ the difference in thermal expansion coefficient between glass and plastic. One approach is to warm the dish on a hot plate and carefully lift up the edge of the coverslip with a sharp razor blade. The approach used by the author is to dip the dish into liquid nitrogen repeatedly—quick dips at first and successively longer dips until the glass cracks and can be removed cautiously with a razor blade or tweezers. As always when working with liquid nitrogen, safety glasses and thermal gloves should be worn. Only a rather small volume of liquid nitrogen is needed and can be decanted from a tank into a small styrofoam box. Tweezers can be used to hold the edge of the dish as it is dipped into the liquid nitrogen. Be careful not to scratch the exposed resin surface with the razor blade as this will damage the thin monolayer of cells. Often, the thin coverslip glass will chip. Care should be taken to remove all the glass as even tiny pieces of glass can damage a diamond knife used for sectioning. 2. Find the photooxidized areas on each dish by comparing the fluorescence and DIC images collected on the light microscope with the light brown cells observed with a dissecting scope (10–20 magnification is usually sufficient). 3. Cut out the region containing the photooxidized cells with a jeweler’s hacksaw after securing the dish on a vise. To keep the top and bottom of the cut-out resin block distinct, use a permanent marker to “color” the top of the block (cells are on the bottom of the dish/block) before cutting. The top of the block will be glued face-down in the next step. 4. Use superglue to secure the cut-out region to an acrylic dummy block that will fit into the ultramicrotome used. It is convenient to use a permanent marker to label the dummy block with the sample information. 5. After mounting the block on an ultramicrotome, trim the block with a razor blade to include only a little more area than the photooxidized
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region. For orientation purposes on the electron microscope, one can make the block face a trapezoidal shape, or notch an edge. 6. For conventional transmission electron microscopy, thin sections of about 70-nm thickness are commonly used. For electron tomography, typical section thickness is 300–500 nm. Because the cells are in a monolayer, trimming the block surface is not advised as it will remove the part of the cells closest to the dish surface. Thus, care must be taken to align the ultramicrotome before cutting sections. Consider using a coated slot grid so that the photooxidized area is not over a grid bar. There is no need to poststain the section because the contrast will come from the label. 7. On the electron microscope, find the photooxidized areas by searching for areas of higher contrast. See supplemental figure 9 of Shu et al. (2011) for examples of the difference in contrast between transfected and untransfected cells.
6. PHOTOOXIDATION PROTOCOL FOR TISSUES Most of the procedure described above for cultured cells applies to slices of tissues from rodents. Up to 200 micron-thick slices have shown good miniSOG labeling throughout the volume. Note that miniSOG has yet to work in plants because the background has been too high. Those parts of the procedure that need modification are described here. 1. Fix the rodent by vascular perfusion first with Ringer’s solution kept in a 37 C water bath to flush out the blood followed by 4% formaldehyde (37 C) made fresh from paraformaldehyde (Electron Microscopy Sciences) in 0.1 M cacodylate buffer. 2. Remove the tissue and place in a 20 mL scintillation vial (Fisher Scientific) in the same fixative on ice for 1 h. Note that glutaraldehyde in combination with paraformaldehyde may increase autofluorescence to the extent that miniSOG fluorescence is obscured making it difficult to find the transfected cells and thus is not added at this step. 3. Slice the tissue using a vibratome (Leica) cooled in an ice bath into 80–200 micron-thick slices. Collect enough slices to replicate the findings and place in the vial. 4. Identify the transfected cells of interest using confocal microscopy after transferring each section to a MatTek dish. Keep the dish at 4 C. 5. Fix the slices with 2% glutaraldehyde for 30 min on ice. 6. Wash the slices in 5 2 min in ice-cold 0.1 M sodium cacodylate buffer.
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7. Block and then photooxidize as described above. 8. Follow the CLEM processing described above except flat-embed the slices in Durcupan resin between two liquid release-agent coated glass slides (Electron Microscopy Sciences). The release agent will allow the glass slides to be separated from the slices after taking out of the oven. Label both slides with the sample information using a permanent marker. 9. Remove one of the glass slides using a razor blade. 10. Find the photooxidized areas that will appear as light brown cells using a dissecting light microscopy. 11. Cut out each photooxidized area with a razor blade. 12. Mount on a dummy block, section and image as above.
7. EXAMPLE OF miniSOG USE WITH MCU An example of the use of miniSOG to label a mitochondrial protein to answer a biological question is, “where do the N and C termini of the mitochondrial calcium uniporter (MCU) protrude—matrix side or intermembrane space side?” The MCU resides in the mitochondrial inner membrane. The report that both the N- and C-termini of MCU protrude into the mitochondrial matrix (Baughman et al., 2011) was countered by another report that both termini face the opposite way—toward the intermembrane space (De Stefani, Raffaello, Teardo, Szabo, & Rizzuto, 2011). Figure 10.2 shows EM of a 1-nm-thick computational slice through a tomographic volume of a cell transfected with miniSOG-MCU. The dark label, indicating MCU location, was found exclusively in the intermembrane space, suggesting that both the N- and C-termini of MCU face outward. Moreover, the labeling was not homogeneous, but instead revealed subdomains of MCU concentration (also see Supplementary Movies 1 and 2 on http://dx.doi.org/10.1016/B978-0-12-801415-8.00010-2). This clarification of MCU topology will guide the search for MCU-interacting partners and proteins that may regulate its function.
8. CONCLUSIONS AND FUTURE WORK miniSOG can be used as a genetic tag method to localize mitochondrial proteins at the resolution provided by electron microscopy while preserving the ultrastructural landmarks that cell biologists use to navigate through organelles, cells, tissues, and organisms. The following bulleted
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Figure 10.2 A 1-nm-thick computational slice through a tomographic volume of a cell transfected with miniSOG-MCU. A cluster of about 20 mitochondria was found close to the nucleus (N). Their membranes are faint but discernible, including the cristae membranes. The dark label, indicating MCU location, was found exclusively in the intermembrane space between the mitochondrial outer membrane and inner boundary membrane. Of interest, the labeling was not homogeneous, but instead revealed subdomains of MCU concentration.
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items provide a glimpse of the immediate future for extensions of miniSOG capability. • Although currently the method is limited to one tag at a time, progress is being made on multi-“color” miniSOG tags by chelating lanthanides, such as cerium and lanthanum, to DAB that allow for elemental mapping of multiple labeled proteins using electron microscopy. • Work is underway to develop viral systems to introduce the miniSOG construct as well as associated fluorescence markers using stereotaxic injection of a lentivirus or the broadly used Cre-lox system. • miniSOG can be combined with cryo-preservation such as high pressure freezing to better preserve cell and tissue ultrastructure. • miniSOG has now been incorporated into TimeSTAMP[YFP] so that both fluorescence and electron microscopy marker detection can be drug-controlled (Butko et al., 2012). TimeSTAMP[YFPminiSOG] thus allows for visualizing protein expression as well as protein turnover by CLEM. This technique has yet to be performed on mitochondrial proteins. • Chromophore-assisted light inactivation (CALI) uses reactive oxygen species (Wojtovich & Foster, 2014) to selectively inactivate a protein of interest by placing a chromophore in the proximity of the protein. The reactive oxygen species generated by the chromophore during illumination oxidize nearby susceptible peptide residues (cysteine, histidine,
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methionine, tryptophan, and tyrosine), greatly reducing the protein’s function. miniSOG, acting as the chromophore, has been used successfully for CALI by genetic fusion with the targeted proteins (Lin et al., 2013). Again, this technique has yet to be performed on mitochondrial proteins. Because miniSOG has considerably longer average fluorescence lifetime compared to the GFP family, it would be useful for F€ orster resonance energy transfer analysis of protein–protein interactions. Finally, exploration has been initiated to combine miniSOG with an even newer genetic tag used for electron microscopy, APEX (Martell et al., 2012). Both use osmication of DAB polymers to contrast the labeled protein. The principle difference is that miniSOG uses a photoreaction, and APEX uses an enzymatic reaction.
ACKNOWLEDGMENTS Tom Deerinck provided the semi-thick section used for electron tomography of miniSOGMCU. This work was partially supported by National Institute of Health grants P01 DK54441, 5P41RR004050, and P41GM103412-24.
REFERENCES Baughman, J. M., Perocchi, F., Girgis, H. S., Plovanich, M., Belcher-Timme, C. A., Sancak, Y., et al. (2011). Integrative genomics identifies MCU as an essential component of the mitochondrial calcium uniporter. Nature, 476, 341–345. Butko, M. T., Yang, J., Geng, Y., Kim, H. J., Jeon, N. L., Shu, X., et al. (2012). Fluorescent and photo-oxidizing TimeSTAMP tags track protein fates in light and electron microscopy. Nature Neuroscience, 15, 1742–1751. De Stefani, D., Raffaello, A., Teardo, E., Szabo, I., & Rizzuto, R. (2011). A forty-kilodalton protein of the inner membrane is the mitochondrial calcium uniporter. Nature, 476, 336–340. Ellisman, M. H., Deerinck, T. J., Shu, X., & Sosinsky, G. E. (2012). Picking faces out of a crowd: Genetic labels for identification of proteins in correlated light and electron microscopy imaging. Methods in Cell Biology, 111, 139–155. Lin, J. Y., Sann, S. B., Zhou, K., Nabavi, S., Proulx, C. D., Malinow, R., et al. (2013). Optogenetic inhibition of synaptic release with chromophore-assisted light inactivation (CALI). Neuron, 79, 241–253. List, N. H., Pimenta, F. M., Holmegaard, L., Jensen, R. L., Etzerodt, M., Schwabe, T., et al. (2014). Effect of chromophore encapsulation on linear and nonlinear optical properties: The case of “miniSOG”, a protein-encased flavin. Physical Chemistry Chemical Physics, 16, 9950–9959. Ludwig, A., Howard, G., Mendoza-Topaz, C., Deerinck, T., Mackey, M., Sandin, S., et al. (2013). Molecular composition and ultrastructure of the caveolar coat complex. PLoS Biology, 11, e1001640. Martell, J. D., Deerinck, T. J., Sancak, Y., Poulos, T. L., Mootha, V. K., Sosinsky, G. E., et al. (2012). Engineered ascorbate peroxidase as a genetically encoded reporter for electron microscopy. Nature Biotechnology, 30, 1143–1148.
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Pimenta, F. M., Jensen, R. L., Breitenbach, T., Etzerodt, M., & Ogilby, P. R. (2013). Oxygen-dependent photochemistry and photophysics of “MiniSOG”, a proteinencased flavin. Photochemistry & Photobiology, 89, 1116–1126. Ruiz-Gonzalez, R., Cortajarena, A. L., Mejias, S. H., Agut, M., Nonell, S., & Flors, C. (2013). Singlet oxygen generation by the genetically encoded tag miniSOG. Journal of the American Chemical Society, 135, 9564–9567. Shu, X., Lev-Ram, V., Deerinck, T. J., Qi, Y., Ramko, E. B., Davidson, M. W., et al. (2011). A genetically encoded tag for correlated light and electron microscopy of intact cells, tissues, and organisms. PLoS Biology, 9, e1001041. Wingen, M., Potzkei, J., Endres, S., Casini, G., Rupprecht, C., Fahlke, C., et al. (2014). The photophysics of LOV-based fluorescent proteins—New tools for cell biology. Photochemical & Photobiological Sciences, 13, 875–883, PMID: 24500379. Wojtovich, A. P., & Foster, T. H. (2014). Optogenetic control of ROS production. Redox Biology, 2, 368–376.
The Use of miniSOG in the Localization of Mitochondrial Proteins Guy A. Perkins1 National Center for Microscopy and Imaging Research, University of California, San Diego, California, USA 1 Corresponding author: e-mail address: [email protected]
Contents 1. 2. 3. 4. 5.
Introduction Requirements for CLEM Labeling miniSOG Features Resolution Photooxidation Protocol for a Monolayer of Cultured Cells 5.1 Chimera construction and transfection notes 6. Photooxidation Protocol for Tissues 7. Example of miniSOG Use with MCU 8. Conclusions and Future Work Acknowledgments References
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Abstract In this chapter, we provide details along with considerations and future directions for the use of miniSOG (for mini Singlet Oxygen Generator), a versatile label for correlated light and electron microscopy of genetically tagged proteins in cells, tissues, and organisms. This new visualizable genetic tag improves the ability of biologists to locate specific proteins at nanoscale resolution and to see these tagged proteins in the environment of structural landmarks that we are used to navigating by, such as mitochondrial membranes and compartments. miniSOG provides high-quality ultrastructural preservation and permits three-dimensional protein localization via electron tomography or serial section block-face scanning electron microscopy. miniSOG is now doing for electron microscopy what the family of green fluorescent protein did for fluorescence microscopy.
Methods in Enzymology, Volume 547 ISSN 0076-6879 http://dx.doi.org/10.1016/B978-0-12-801415-8.00010-2
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2014 Elsevier Inc. All rights reserved.
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1. INTRODUCTION CLEM—correlated light and electron microscopy (the same area examined by both microscopy techniques) or correlative light and electron microscopy (the same specimen, but not the same area)—is an expanding tool for dissecting the protein landscape useful for providing clues to cell and tissue function at high spatial resolution (Ellisman, Deerinck, Shu, & Sosinsky, 2012). Each imaging modality provides unique information, and the combination of light and electron microscopies can contribute to a better understanding of the spatiotemporal patterns of protein expression, trafficking, binding partners, and function. Fluorescence light microscopy provides the dynamics and options for multiprotein labeling, and electron microscopy provides the increased resolution and cellular context. The use of genetically appended tags to specific proteins to be able to localize them in their complex cellular milieu is growing as new tags have been developed.
2. REQUIREMENTS FOR CLEM LABELING Certain requirements need to be met for a genetic tag to be useful as a CLEM label. First and foremost, the genetic sequence of the protein of interest must be known. Second, a genetic tag must be nontoxic to the cell in which it is expressed. It should be active in the physiological environment where expressed. In addition to the exhibiting fluorescence, it should either independently or with additional ligands or cofactors create an electrondense label for CLEM imaging. A genetic tag has the additional attractive property that it is stoichiometric to its target protein. This quality can be important for quantification of low copy number proteins within cells. As with any exogenously expressed protein, it is important to conduct control experiments to ensure that recombinant proteins are expressed and localized the same as endogenously expressed proteins. A commonly used control is comparison to immunolabeled native systems.
3. miniSOG FEATURES Mini Singlet Oxygen Generator (miniSOG) is one such recently engineered genetic tag. It is a 106-amino acid green fluorescent flavoprotein generated from Arabidopsis phototropin2 that efficiently generates singlet oxygen when illuminated with blue light (Shu et al., 2011). To generate
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Figure 10.1 When excited by blue light (dark gray in print version), miniSOG produces singlet oxygen, a reactive and short-lived excited state of O2. This singlet oxygen quickly oxidizes and polymerizes diaminobenzidine (DAB) into a localized osmiophilic density. Addition of osmium tetroxide to the polymerized DAB produces an electron-opaque stain that then allows nanometer-scale imaging of the tagged protein by electron microscopy.
singlet oxygen, miniSOG requires a flavin mononucleotide (FMN) cofactor. FMN, derived from the vitamin riboflavin, is endogenous to cells and is ubiquitous. It is indispensable in mitochondrial electron transport, fatty acid oxidation, and vitamin metabolism. Singlet oxygen, a reactive and short-lived excited state of O2, quickly oxidizes and polymerizes diaminobenzidine (DAB) into a localized osmiophilic mass, which osmicated density is clearly seen in the electron microscope (Fig. 10.1). Diffusion of this dense reaction product is minimized by extensive chemical cross-linking by glutaraldehyde prior to the generation of the reaction product. Because miniSOG is relatively small, about half the size of GFP, the protein tagging is less likely to influence protein targeting to the right location than larger genetic tags. Because miniSOG is genetically fused to the protein of interest and all other components (DAB, light, O2, and OsO4) are small and readily permeate well-fixed tissue, there is no conflict between labeling efficiency and established methods for excellent structural preservation. Moreover, miniSOG can photooxidize DAB within relatively thick slices of tissue, expanding its applicability. Indeed, miniSOG fusions have been expressed in adult mice using in utero electroporation. miniSOG causes no obvious cellular toxicity in the absence of light. Table 10.1 summarizes the properties of the miniSOG genetic label. The original report of a high quantum yield of singlet oxygen (0.47) has recently been challenged in two reports (Pimenta, Jensen, Breitenbach, Etzerodt, & Ogilby, 2013; Ruiz-Gonzalez, Cortajarena, Mejias, Agut, Nonell and Flors,
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Table 10.1 miniSOG Properties 1
Size (kDa)
Excitation (nm)
Emission (nm)
O2 Quantum yield
Fluorescent quantum yield
Brightness (M21 cm21)
13.9
447/473 (shoulder)
500/528 (shoulder)
0.47 (0.03a)
0.37
5820
a Pimenta et al. (2013) and Ruiz-Gonzalez et al. (2013) report a lower quantum yield of 0.03. Adapted from Shu et al. (2011) and table 1 of Wingen et al. (2014). Additional spectroscopic information is provided by List et al. (2014).
2013) that measured a quantum yield of 0.03 for miniSOG. The difference was attributed to the finding that miniSOG oxidizes ADPA, the standard singlet oxygen sensor used by both singlet oxygen-dependent and -independent processes. Interestingly, the low intrinsic quantum yield is compensated by a photoinduced transformation of miniSOG by cumulative irradiation that increases its quantum yield about 10-fold, which corroborates its utility for CLEM. miniSOG fusions have been made for many well characterized proteins and have been found to localize correctly in cultured cells, tissues, and intact organisms enabling CLEM without the need for exogenous ligands, probes, or destructive permeabilizing detergents. Therefore, miniSOG can be used for high-resolution 3D electron microscopy modalities, such as electron tomography and serial block-face scanning electron microscopy. This combination of high-resolution protein labeling and 3D electron microscopy now permits scientists to pursue answers to questions previously impossible to ask, such as does a particular mitochondrial protein localizes to the crista junction? Unlike particulate markers such as protein-conjugated immunogold or quantum dot labels, miniSOG does not just decorate the targeted protein but also is similar to a negative stain in that it outlines the protein (Ludwig et al., 2013). miniSOG can be used alone or in combination with different fluorescent proteins as the optical reporter for the tagged protein. Because miniSOG photobleaches rather quickly it thus may be necessary to fuse it to a second, nonoverlapping fluorescent genetic marker such as mCherry, tdTomato, or mKO2.
4. RESOLUTION The ultimate resolution achievable by miniSOG labeling remains to be determined. However, the diffusion of the reacted DAB product away from the miniSOG tag has been has been shown to be minimal, thus
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allowing proteins to be localized within a few nanometer precision (Ludwig et al., 2013). Using line scans across images of miniSOG-labeled caveolar coat protein, this group found that the polymerized and osmicated DAB electron density clearly resolved periodic density changes of about 10 nm. To accurately determine the resolution, one could express miniSOG on a subunit of a periodic solved structure, such as an isoform of actin or tubulin or an actin-associated protein. High-resolution electron tomography would then be performed. Subvolume averaging of tomographic reconstructions would define the molecular envelope of the tagged protein, which would then allow for the determination of the location of the osmium DAB density relative to the protein.
5. PHOTOOXIDATION PROTOCOL FOR A MONOLAYER OF CULTURED CELLS 5.1. Chimera construction and transfection notes Fusion constructions and transfections that work for GFP-family labeled proteins usually work for miniSOG fusions. Chemical transformation or lentiviral infections are often the most convenient, including stereotaxic injection of a lentivirus into a specific organ. However, electroporation is the method of choice for large plasmids. miniSOG fusions can be cloned into the pcDNA3.1 vector (Invitrogen). miniSOG or chimera cDNAs can be transfected into the cell line of choice using the Fugene (Promega) kit. Successful transfection of neurons has been achieved using Amaxa electroporation (Lonza AG, Germany). For rodents, miniSOG fusion constructs can be delivered into the lateral ventricle of embryos by in utero electroporation. It is recommended that two people be involved in the experiment to share the making of solutions, and performing the incubations and washes in order to not leave the samples in the various solutions too long. When removing solutions, it is important not to touch the central circle of the dish where the glass coverslip is as this will be the region imaged with CLEM. When adding solutions, it is important not to direct the stream of liquid on this central portion of the dish, but rather to the side of the dish so as not to disturb the cells. Note that many of the chemicals used are toxic, and so appropriate lab safety procedures should be followed. 1. Fix transfected cells cultured on glass bottom culture dishes (P35G-014-C, MatTek Corp., Ashland, MA) with 2% glutaraldehyde + 2 mM CaCl22H2O in 0.1 M sodium cacodylate buffer (pH 7.4) for 60 min.
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Warm fixative to 37 C. After removing the growth medium from these samples by either pouring off or pipetting off around the dish edge with a plastic disposable pipet, add the warmed fixative. Note that a glass pipet should not be used because microshards of glass may flake off and if embedded in the plastic resin may damage a diamond knife used for sectioning. Let the samples cool to room temperature for 5 min. Place the samples on ice for the last 55 min. Note that all aqueous solutions should be made with double-distilled water (ddH2O). From this step forward, it is important to keep the samples cold at all times until the 100% ethanol step below. Make up the cacodylate washing buffer and blocking solution (next 2 steps). Get them cooling on ice. It is wise to look one to two steps ahead to have chilled solutions ready when needed. 2. Wash samples with ice-cold 0.1 M sodium cacodylate buffer (pH 7.4) five times for 2 min each (5 2 min). Keep samples on ice. It is important that each sample not dry out by being uncovered by solution for more than a few seconds. So, when washing or replacing solutions, add the new solution quickly after removing the previous and do this for each sample before going to the next sample, i.e., do not remove the solution from all the samples before adding the new solution to all the samples. 3. Photoconversion block: Block samples for 15–20 min with ice-cold blocking solution consisting of 20 mM glycine, 10 mM potassium cyanide, and 10 mM aminotriazole (catalase inhibitor) in 0.1 M sodium cacodylate buffer. Right before use, add hydrogen peroxide: 1 μL per 25 mL of blocking solution. Keep samples on ice. 4. Wash samples with ice-cold 0.1 M cacodylate buffer 5 2 min. Keep samples on ice. Optional: Additional blocking step with mersalyl acid to reduce background nonspecific labeling of mitochondria. Note that the mitochondrial electron transport chain can generate singlet oxygen. Mersalyl acid is a mercury compound that poisons the ETC to block this singlet oxygen production. a. Block samples in 5–10 mM mersalyl acid (Sigma Cat. No. M9784-1G) for 30 min on ice. b. To make 10 mL of a 5 mM solution Dissolve 24 mg (50 μM) of the mersalyl acid in 50 μL of a 1 M NaOH stock solution. Vortex until almost all the mersalyl acid has been dissolved. Add 6.7 mL ddH2O + 3.3 mL of 0.3 M
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cacodylate buffer. Add 35 μL of a 1 M HCl stock solution. Filter with a 0.22 μm Millipore filter attached to a syringe. Cool this solution on ice before adding to the samples. 5. Wash samples in ice-cold 0.1 M cacodylate buffer 5 2 min. Keep samples on ice. 6. Make the DAB solution: DAB (free base—MW 214.27) (Cat. No. D8001-10G Sigma) Dissolve 0.005 mg free base DAB in 1 mL 0.1 M HCl (final concentration 25 mM stock solution) by vortexing until dissolved (may take a couple of minutes). Add 9 mL of 0.1 M cacodylate buffer. Filter solution into a new 20 mL scintillation vial (Fisher Scientific) with a 0.22 μm Millipore filter attached to a syringe. Photooxidation steps 1. Use an inverted confocal or inverted fluorescence microscope to identify transfected cells and for correlative light microscopic imaging. An inverted microscope ensures direct open access to the DAB solution. Set the microscope stage temperature to 4 C. 2. Take initial picture with minimal exposure to avoid photobleaching of the target area on the sample dish using both fluorescence and DIC. 3. Remove buffer from the dish and add enough DAB solution to cover the bottom of the dish. Let sit for 5 min. 4. Take off the dish lid. Gently blow oxygen continuously from an oxygen tank onto the sample over the top of the liquid surface. Alternately, the DAB solution on ice can be bubbled with oxygen and the solution in the dish refreshed every 1–2 min. 5. Illuminate the target area using a standard FITC filter set (EX470/40, DM510, BA520) with light from a 150 W xenon lamp to photooxidize until the sample turns a light brown as monitored by transmitted light. It is important to know that photooxidation is fast (typically 2–6 min) depending on how concentrated the labeled protein is and how accessible the miniSOG is in the folded protein. It is recommended that optimal timing to produce a well-visualized reaction product, but not to overreact, be determined by screening a small series of time points. If the illumination was too long, background density is observed in the electron microscope. Care should be taken not to illuminate with too intense beam as this may cause photodamage to cells. A setting of 30% power is typically used for the xenon lamp. Mitochondria are particularly sensitive and the damage most often seen is swollen matrix or matrix with regions devoid of cristae.
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6. Take final fluorescence and DIC pictures after photooxidation. Go to the next area in the same dish and repeat, usually three to four areas per dish. 7. Remove the sample from the microscope and wash it in ice-cold 0.1 M cacodylate buffer 5 2 min. Keep samples in buffer on ice until done with all plates. 8. Incubate samples in 1% osmium tetroxide (Electron Microscopy Sciences) in 0.1 M cacodylate buffer 30 min on ice. Work in the hood because osmium tetroxide is toxic and volatile. Electron microscope sample preparation steps 1. Wash samples in ice-cold ddH2O 3 2 min. Keep samples on ice. 2. Stain en bloc with 2% aqueous uranyl acetate (Ted Pella Inc.) 1 h on ice. If needed, this step can go overnight. Simply place the dishes in a refrigerator. Note that the first time doing the experiment it is recommended to have one plate sans uranyl acetate as it may mask the true signal, which is a higher concentration (contrast) of osmium bound to the DAB, especially relevant if the protein is sparse. 3. Dehydrate the samples in an ice-cold graded ethanol series (20%, 50%, 70%, and 90%) 2 min each. 4. Further dehydrate in 100% anhydrous ethanol 3 2 min at room temperature. Remove as much of the liquid as possible from each dehydration by pipet. Make sure that no water gets inside the dish during these dehydrations, and, of course in subsequent steps. Keep the lid on the dish to minimize water vapor on the samples. Try not to breathe over the dishes to minimize water vapor. Remove the ice bucket from the vicinity of the dishes to avoid water condensation close to the samples. From here to the end of the procedure, water is the enemy. 5. Infiltrate the samples in Durcupan ACM resin (Electron Microscopy Sciences) using 1:1 anhydrous ethanol:resin for 30 min at room temperature. The ethanol acts as a vehicle to aid the viscous resin to infiltrate into the cells. The four different bottles used are labeled A (or A/M), B, C, and D, respectively. The ratios of the Durcupan components should be 11.4 mL A:10 mL B:0.3 mL C:0.1 mL D. Because certain components are viscous, measure 11.4 g of A and 10 g B on a scale. For C and D, use 1 mL syringes to draw the solution from the bottle. Component C is smelly, so work in the hood. 6. Infiltrate the samples in 100% Durcupan ACM resin 3 1 h with freshly made resin each time because the resin hardens with time and this hardening impedes infiltration. Remove as much of the previous resin as
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possible taking care not to touch the central circle of the dish, where the glass cover slip is glued. At this point, ethanol is the enemy as even a little residual ethanol in the resin will make the samples soft and thus hard to section. 7. Polymerize the samples in an oven set to 60–70 C for 48 h. After the samples have cooled to room temperature, they should be “rock-hard,” important for good sectioning. Electron microscopy 1. Remove the glass coverslip from the bottom of the dish. It is usually difficult to remove this coverslip, and thus the strategy is to employ the difference in thermal expansion coefficient between glass and plastic. One approach is to warm the dish on a hot plate and carefully lift up the edge of the coverslip with a sharp razor blade. The approach used by the author is to dip the dish into liquid nitrogen repeatedly—quick dips at first and successively longer dips until the glass cracks and can be removed cautiously with a razor blade or tweezers. As always when working with liquid nitrogen, safety glasses and thermal gloves should be worn. Only a rather small volume of liquid nitrogen is needed and can be decanted from a tank into a small styrofoam box. Tweezers can be used to hold the edge of the dish as it is dipped into the liquid nitrogen. Be careful not to scratch the exposed resin surface with the razor blade as this will damage the thin monolayer of cells. Often, the thin coverslip glass will chip. Care should be taken to remove all the glass as even tiny pieces of glass can damage a diamond knife used for sectioning. 2. Find the photooxidized areas on each dish by comparing the fluorescence and DIC images collected on the light microscope with the light brown cells observed with a dissecting scope (10–20 magnification is usually sufficient). 3. Cut out the region containing the photooxidized cells with a jeweler’s hacksaw after securing the dish on a vise. To keep the top and bottom of the cut-out resin block distinct, use a permanent marker to “color” the top of the block (cells are on the bottom of the dish/block) before cutting. The top of the block will be glued face-down in the next step. 4. Use superglue to secure the cut-out region to an acrylic dummy block that will fit into the ultramicrotome used. It is convenient to use a permanent marker to label the dummy block with the sample information. 5. After mounting the block on an ultramicrotome, trim the block with a razor blade to include only a little more area than the photooxidized
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region. For orientation purposes on the electron microscope, one can make the block face a trapezoidal shape, or notch an edge. 6. For conventional transmission electron microscopy, thin sections of about 70-nm thickness are commonly used. For electron tomography, typical section thickness is 300–500 nm. Because the cells are in a monolayer, trimming the block surface is not advised as it will remove the part of the cells closest to the dish surface. Thus, care must be taken to align the ultramicrotome before cutting sections. Consider using a coated slot grid so that the photooxidized area is not over a grid bar. There is no need to poststain the section because the contrast will come from the label. 7. On the electron microscope, find the photooxidized areas by searching for areas of higher contrast. See supplemental figure 9 of Shu et al. (2011) for examples of the difference in contrast between transfected and untransfected cells.
6. PHOTOOXIDATION PROTOCOL FOR TISSUES Most of the procedure described above for cultured cells applies to slices of tissues from rodents. Up to 200 micron-thick slices have shown good miniSOG labeling throughout the volume. Note that miniSOG has yet to work in plants because the background has been too high. Those parts of the procedure that need modification are described here. 1. Fix the rodent by vascular perfusion first with Ringer’s solution kept in a 37 C water bath to flush out the blood followed by 4% formaldehyde (37 C) made fresh from paraformaldehyde (Electron Microscopy Sciences) in 0.1 M cacodylate buffer. 2. Remove the tissue and place in a 20 mL scintillation vial (Fisher Scientific) in the same fixative on ice for 1 h. Note that glutaraldehyde in combination with paraformaldehyde may increase autofluorescence to the extent that miniSOG fluorescence is obscured making it difficult to find the transfected cells and thus is not added at this step. 3. Slice the tissue using a vibratome (Leica) cooled in an ice bath into 80–200 micron-thick slices. Collect enough slices to replicate the findings and place in the vial. 4. Identify the transfected cells of interest using confocal microscopy after transferring each section to a MatTek dish. Keep the dish at 4 C. 5. Fix the slices with 2% glutaraldehyde for 30 min on ice. 6. Wash the slices in 5 2 min in ice-cold 0.1 M sodium cacodylate buffer.
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7. Block and then photooxidize as described above. 8. Follow the CLEM processing described above except flat-embed the slices in Durcupan resin between two liquid release-agent coated glass slides (Electron Microscopy Sciences). The release agent will allow the glass slides to be separated from the slices after taking out of the oven. Label both slides with the sample information using a permanent marker. 9. Remove one of the glass slides using a razor blade. 10. Find the photooxidized areas that will appear as light brown cells using a dissecting light microscopy. 11. Cut out each photooxidized area with a razor blade. 12. Mount on a dummy block, section and image as above.
7. EXAMPLE OF miniSOG USE WITH MCU An example of the use of miniSOG to label a mitochondrial protein to answer a biological question is, “where do the N and C termini of the mitochondrial calcium uniporter (MCU) protrude—matrix side or intermembrane space side?” The MCU resides in the mitochondrial inner membrane. The report that both the N- and C-termini of MCU protrude into the mitochondrial matrix (Baughman et al., 2011) was countered by another report that both termini face the opposite way—toward the intermembrane space (De Stefani, Raffaello, Teardo, Szabo, & Rizzuto, 2011). Figure 10.2 shows EM of a 1-nm-thick computational slice through a tomographic volume of a cell transfected with miniSOG-MCU. The dark label, indicating MCU location, was found exclusively in the intermembrane space, suggesting that both the N- and C-termini of MCU face outward. Moreover, the labeling was not homogeneous, but instead revealed subdomains of MCU concentration (also see Supplementary Movies 1 and 2 on http://dx.doi.org/10.1016/B978-0-12-801415-8.00010-2). This clarification of MCU topology will guide the search for MCU-interacting partners and proteins that may regulate its function.
8. CONCLUSIONS AND FUTURE WORK miniSOG can be used as a genetic tag method to localize mitochondrial proteins at the resolution provided by electron microscopy while preserving the ultrastructural landmarks that cell biologists use to navigate through organelles, cells, tissues, and organisms. The following bulleted
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Figure 10.2 A 1-nm-thick computational slice through a tomographic volume of a cell transfected with miniSOG-MCU. A cluster of about 20 mitochondria was found close to the nucleus (N). Their membranes are faint but discernible, including the cristae membranes. The dark label, indicating MCU location, was found exclusively in the intermembrane space between the mitochondrial outer membrane and inner boundary membrane. Of interest, the labeling was not homogeneous, but instead revealed subdomains of MCU concentration.
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items provide a glimpse of the immediate future for extensions of miniSOG capability. • Although currently the method is limited to one tag at a time, progress is being made on multi-“color” miniSOG tags by chelating lanthanides, such as cerium and lanthanum, to DAB that allow for elemental mapping of multiple labeled proteins using electron microscopy. • Work is underway to develop viral systems to introduce the miniSOG construct as well as associated fluorescence markers using stereotaxic injection of a lentivirus or the broadly used Cre-lox system. • miniSOG can be combined with cryo-preservation such as high pressure freezing to better preserve cell and tissue ultrastructure. • miniSOG has now been incorporated into TimeSTAMP[YFP] so that both fluorescence and electron microscopy marker detection can be drug-controlled (Butko et al., 2012). TimeSTAMP[YFPminiSOG] thus allows for visualizing protein expression as well as protein turnover by CLEM. This technique has yet to be performed on mitochondrial proteins. • Chromophore-assisted light inactivation (CALI) uses reactive oxygen species (Wojtovich & Foster, 2014) to selectively inactivate a protein of interest by placing a chromophore in the proximity of the protein. The reactive oxygen species generated by the chromophore during illumination oxidize nearby susceptible peptide residues (cysteine, histidine,
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methionine, tryptophan, and tyrosine), greatly reducing the protein’s function. miniSOG, acting as the chromophore, has been used successfully for CALI by genetic fusion with the targeted proteins (Lin et al., 2013). Again, this technique has yet to be performed on mitochondrial proteins. Because miniSOG has considerably longer average fluorescence lifetime compared to the GFP family, it would be useful for F€ orster resonance energy transfer analysis of protein–protein interactions. Finally, exploration has been initiated to combine miniSOG with an even newer genetic tag used for electron microscopy, APEX (Martell et al., 2012). Both use osmication of DAB polymers to contrast the labeled protein. The principle difference is that miniSOG uses a photoreaction, and APEX uses an enzymatic reaction.
ACKNOWLEDGMENTS Tom Deerinck provided the semi-thick section used for electron tomography of miniSOGMCU. This work was partially supported by National Institute of Health grants P01 DK54441, 5P41RR004050, and P41GM103412-24.
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Pimenta, F. M., Jensen, R. L., Breitenbach, T., Etzerodt, M., & Ogilby, P. R. (2013). Oxygen-dependent photochemistry and photophysics of “MiniSOG”, a proteinencased flavin. Photochemistry & Photobiology, 89, 1116–1126. Ruiz-Gonzalez, R., Cortajarena, A. L., Mejias, S. H., Agut, M., Nonell, S., & Flors, C. (2013). Singlet oxygen generation by the genetically encoded tag miniSOG. Journal of the American Chemical Society, 135, 9564–9567. Shu, X., Lev-Ram, V., Deerinck, T. J., Qi, Y., Ramko, E. B., Davidson, M. W., et al. (2011). A genetically encoded tag for correlated light and electron microscopy of intact cells, tissues, and organisms. PLoS Biology, 9, e1001041. Wingen, M., Potzkei, J., Endres, S., Casini, G., Rupprecht, C., Fahlke, C., et al. (2014). The photophysics of LOV-based fluorescent proteins—New tools for cell biology. Photochemical & Photobiological Sciences, 13, 875–883, PMID: 24500379. Wojtovich, A. P., & Foster, T. H. (2014). Optogenetic control of ROS production. Redox Biology, 2, 368–376.