Visualization of miniSOG Tagged DNA Repair Proteins in Combination with Electron Spectroscopic Imaging (ESI)

Podsumowanie

Electron spectroscopic imaging can image and distinguish nucleic acid from protein at nanometer resolution. It can be combined with the miniSOG system, which is able to specifically label tagged proteins in transmission electron microscopy samples. We illustrate the use of these technologies using double-strand break repair foci as an example.

Streszczenie

The limits to optical resolution and the challenge of identifying specific protein populations in transmission electron microscopy have been obstacles in cell biology. Many phenomena cannot be explained by in vitro analysis in simplified systems and need additional structural information in situ, particularly in the range between 1 nm and 0.1 µm, in order to be fully understood. Here, electron spectroscopic imaging, a transmission electron microscopy technique that allows simultaneous mapping of the distribution of proteins and nucleic acids, and an expression tag, miniSOG, are combined to study the structure and organization of DNA double-strand break repair foci.

Wprowadzenie

Despite the significant advancements in light microscopy over recent years¹, cell-biologists still suffer from a gap in resolution. This limits understanding of the structure-function relationships in fundamental cellular processes that involve the coordinated interplay between macromolecular complexes (e.g., in chromatin remodelling, DNA repair, RNA transcription and DNA replication). Although transmission electron microscopes (TEM)s provide the required resolution, it has been challenging to define these processes structurally because of the inability to label specific proteins while also being able to determine the biochemical composition of the visualized structures. In the absence of internal membranes to help differentiate nuclear structures, the nucleus has been particularly challenging. Electron spectroscopic imaging (ESI) solves some of these limitations by allowing the simultaneous detection and differentiation of DNA, RNA, and protein-based nuclear structures^2-5.

Electron spectroscopic imaging:

In order to map elemental distributions at high sensitivity and resolution in the electron microscope, one can use an imaging spectrometer that selects electrons that have been inelastically scattered through interactions with inner shell electrons of an element in the specimen⁶. Because element-specific amounts of energy are lost as a consequence of ionization of atoms in the specimen, these electrons can be separated and visualized using a spectrometer that is attached to the electron microscope. Thus, analysis of the spectrum of the electrons that have interacted with the specimen reveals qualitative and quantitative information about the elemental composition of the sample⁷. Electrons that do not lose energy when passing through the specimen are found in the “zero-loss peak” of the electron energy loss spectrum. The abundance of these electrons is related to the mass, density and thickness of the specimen and is comprised of electrons that pass through the specimen without colliding with the specimen or losing energy during passage through the specimen. This information can be useful for absolute quantification of the numbers of atoms of a specific element present in the specimen⁸.

Since biological samples consist mostly of light elements that poorly deflect the electrons in the incident beam of the TEM, staining methods using heavy metal salts have to be applied in order to generate contrast in the sample. The lack of specificity of most of these contrasting agents and the inability to visualize more than one stain where specificity is possible has limited the value of conventional electron microscopy in the study of the nucleus. ESI has significant advantages over conventional TEM, particularly for the study of structures of the cell nucleus. It is possible to exploit the phosphorus-rich nature of DNA- and RNA-containing macromolecular complexes to distinguish nucleoprotein complexes from protein complexes and to resolve different nucleoprotein complexes based on their density of nucleic acids. The remaining biological material can be imaged based on its abundance of nitrogen. Mapping just these two elements and analysis of their distribution and relative abundance within different anatomical structures provides us with a lot of information about the nucleus. For example, it is easy to identify chromatin and the ribosomes in the map representing phosphorus abundance. The interchromatin space, nuclear pore complexes, and nuclear bodies, on the other hand, can be easily detected in the nitrogen map image.

Mini singlet oxygen generator system (miniSOG)

While ESI represents a powerful technique to study in situ chromatin structure because it takes advantage of the characteristic ratios in elemental composition between phosphorus and nitrogen, the elemental composition cannot normally be used to discriminate between different populations of protein complexes. Antibodies labeled with small gold particles in the nanometer range have been widely used to map the location of individual molecules. Since the gold particle is usually attached to a secondary antibody, it will appear within a circumference of approximately 20 nm around the epitope detected by the primary antibody. In post-embedding samples, antibodies can only detect the epitopes that are exposed at the surface of the sections. While it is possible to prove the presence of an antigen and relate it to a certain anatomical structure of the cell, the information that is obtained is incomplete, since most of the epitopes are obscured by resin. Pre-embedding techniques, which use similar protocols to those used for fluorescence microscopy, allow access to antigens throughout the entire depth of the sample but the permeabilization steps that are required to allow the antibody to penetrate the cell typically require removal of lipid membranes and remove components that cannot be fixed by aldehydes. Moreover, the preferred aldehyde fixative for ultrastructure preservation, glutaraldehyde, commonly destroys epitopes and, consequently, paraformaldehyde is typically required. This is less effective at protein-protein crosslinking. Another disadvantage of antibodies that are labeled with a gold nanoparticle is that gold is a very electron-dense material that creates a strong contrast that can obscure interesting structural details of the sample, which has weaker contrast.

The emergence of the green fluorescent protein (GFP) as an expressed protein tag has transformed the use of fluorescence microscopy to answer questions in cell biology. Tagging a protein with a small fluorescent domain allows mapping of its distribution in vivo without the need for permeabilization procedures that can alter the structure. In order to translate the elegant principle of using protein tags to map protein distributions in a cell to electron microscopy, a system was needed that is able to produce a signal close to the structure of interest and generate contrast in the TEM. Polymerized diaminobenzidine is a stain that is often used in histology to detect antibody binding. This stain is usually deposited using horseradish peroxidase (HRP) conjugated to a secondary antibody. Although the reaction produces a reliable result, HRP is not catalytically active in the cytosol of cells⁹. The reaction products of HRP can also diffuse away from the site of generation so that their resolution is worse than the nanogold method¹⁰. In order to bypass these problems, the FlAsH/ReAsH system was developed¹¹. It consists of recombinant fusion proteins that possess a tetracysteine motif. This motif allows binding of a biarsenic fluorophore. When excited, the bound FlAsH or ReAsH is able to generate the highly reactive singlet oxygen and thus photoconvert diaminobenzidine (DAB) into a polymer that precipitates immediately at the site of the tagged proteins. The DAB polymer can be stained with osmium tetroxide, which is electron dense and thus can be used to map the distribution of the recombinant fusion protein in the TEM.

In 2011, Shu et al.¹⁰ presented the miniSOG system, which consists of a small 106 amino acids fusion tag derived from a flavoprotein of Arabidopsis that is fluorescent and able to create many singlet oxygen radicals when excited with 448 nm blue light. Those singlet oxygen radicals can be used to photo-oxidize diaminobenzidine to form polymers on and near the surface of the tagged protein, which is considerably closer than immunogold labeled antibodies¹¹. While the FlAsH/ReAsH system requires bringing the fluorochrome and the diaminobenzidine into the cells before doing the photoconversion, the miniSOG system only requires diaminobenzidine and light and in addition is about twice as effective at polymerizing diaminobenzidine. Here, miniSOG is employed in combination with ESI in order to map the ultrastructure of DNA repair foci.

DNA repair foci (DRF)

Unrepaired DNA double strand breaks pose a serious threat to the cell since they can lead to translocations and loss of genetic information. In turn, this can lead to senescence, cancer, and cell death. Many proteins that are involved in DNA double strand break repair accumulate in foci that assemble around a DNA double strand break^12-14. Although their function is not known, they represent the site in the nucleus that contains the DNA double strand break and is the site of DNA double strand break repair.

DNA repair foci (DRF) have been characterized by fluorescence microscopy and they serve as biomarkers for DNA damage^12,15. They are massive relative to the size of the double-strand break and were considered relatively homogeneous until recent super-resolution microscopy studies revealed some evidence of sub-compartmentalization of molecules within each focus¹⁶. In order to understand how these sites are organized, it is necessary to visualize all of the underlying biological structures relative to each other. This cannot be achieved by fluorescence microscopy but is possible through electron microscopy^17,18. Here, a method is described that combines electron spectroscopic imaging with the miniSOG method to illustrate the potential of this combined approach to explore the ultrastructure of DNA double-strand break repair.

Protokół

1. Generation of miniSOG Cell-lines

1. Grow U2OS (human osteosarcoma) cells in a 35 mm dish containing 2 ml of low glucose Dulbecco’s modified Eagle’s medium (DMEM) that is supplemented with 10% foetal bovine serum (FBS) at 37 °C in a humidified incubator with 5% CO₂ atmosphere.
2. Transfect the cells when they are 80% confluent by lipofection with purified transfection quality (A269/280 ratio 1.8-2.0) plasmid constructs containing the sequences for miniSOG and mCherry tagged repair proteins according to the protocol of the manufacturer of the transfection reagent¹⁹. The miniSOG expression plasmids can be ordered from the Tsien-Lab: http://www.tsienlab.ucsd.edu/Samples.htm
3. Sort the cells expressing the recombinant protein with the miniSOG and mCherry tag by fluorescence activated cell sorting two days post transfection. Collect cells with intermediate fluorescence intensity in order to avoid cells that are overexpressing²⁰.
4. Culture the positive cells on a 10 cm dish in 10 ml of DMEM medium that is supplemented with 10 % FBS and 0.6 µg/ml of the selection agent G418 (Geneticin).
5. After visible colonies have emerged on the plates, screen for mCherry positive colonies using an inverted fluorescence microscope and draw circles around them (onto the bottom of the plate) with a permanent marker. Use a low magnification air objective lens (e.g., 10x) and pick clones using sterile technique by carefully scratching the colonies off and slowly sucking them into a sterile 1,000 µl pipette tip.
6. Expand the selected colonies and freeze parts of them down using the growth medium described in step 1.1 supplemented with 10% dimethyl sulphoxide. Test the cells for DRF formation by growing them on sterile 18 x 18 mm² cover slips and exposing them to 2 Gy of radiation. The irradiated cells stably expressing a miniSOG mCherry tagged repair protein must show the typical focal pattern characteristic of DSBs when visualized with a fluorescence microscope using a filter-set for imaging mCherry.

2. Cell Culture

1. Culture U2OS cells stably expressing the miniSOG and mCherry tagged repair proteins under the conditions outlined in step 1.1. Grow cells to 80% confluency in a 35 mm diameter glass bottom dish containing 2 ml of DMEM. Make sure that the glue that attaches the cover slip to the plastic and the used plastic is resistant to aqueous and alcoholic solutions and resistant to the resin used!
2. Before conducting the experiment, outline a small area of approximately 1 mm² that contains cells expressing the ectopic proteins by scratching with a sterile diamond pen using a fluorescence microscope to identify the region of interest. Alternatively use a glass bottom dish that contains a coverslip with a pre-etched grid system built into the coverslip.
   NOTE: If using high quality correlative microscopy combining ESI and fluorescence microscopic pictures²¹, make sure that the thickness of the coverslip is compatible with oil immersion objectives. It should have the thickness No. 1.5 (0.16-0.18 mm). Choose an area  close to the center of the dish for the region to be examined in the TEM in order to avoid problems with polymerization of the resin that may occur near the edges (see point 4.10-4.13).

3. DNA Damage Induction

1. Irradiate the cells with gamma radiation, use a radiomimetic drug, or induce the damage by laser micro irradiation on a confocal microscope (e.g., as described in^18,22 or ²³).
2. In this example, irradiate the cells with 2 and 6 Gy of gamma irradiation in a cesium-137 radiation source or laser microirradiate using the 405 nm solid state laser of a confocal microscope after sensitizing the cells for 20 min with 0.5 µg/ml Hoechst.

4. Sample Preparation

1. Fix the cells with 1 ml of 4% paraformaldehyde CAUTION in 0.1 M sodium cacodylate buffer or in 0.1 M phosphate buffer pH 7.4 for 30 min at RT in the dark.
   CAUTION: Paraformaldehyde and sodium cacodylate are toxic! Wear gloves and dispose the toxic substances adequately.
2. Wash the cells twice with 4 ml of 0.1 M sodium cacodylate buffer pH 7.4 or phosphate buffer pH 7.4.
3. Treat the fixed cells with 2 ml of 50 mM glycine, 5 mM aminotriazole and 10 mM potassium cyanide CAUTION in 0.1 M sodium cacodylate buffer or 0.1 M phosphate buffer (pH 7.4) (check the pH!) for 30 min in order to block unreacted aldehyde groups and to suppress the reactive oxygen species generated by heme groups, which can increase the background.
   CAUTION: Potassium cyanide is extremely toxic! Wear gloves, work under a hood and dispose the toxic substances adequately. The reaction is very sensitive to pH so it is important that the pH verified and accurate.
4. Record the area that was selected in step 2.2 in 2D or 3D on an inverted fluorescence microscope, if correlative microscopy is desired.
5. Prepare a 1 mg/ml diaminobenzidine hydrochloride CAUTION solution by placing a 10 mg diaminobenzidine hydrochloride tablet into a 2 ml microcentrifuge tube. Add 980 µl of distilled H₂O and 20 µl of concentrated hydrochloric acid (11.65 M) and mix until the solution is light brown and translucent. Dilute this solution in 0.1 M phosphate or 0.1 M sodium cacodylate buffer to a final concentration of 1 mg/ml while adjusting the pH with sodium hydroxide to a pH between 7.0 and 7.6 (pH 7.4 is recommended).
   CAUTION: Diaminobenzidine is toxic! Wear gloves and dispose the toxic substances adequately.
6. For photooxidation, replace the buffer with 2 ml of a 1 mg/ml diaminobenzidine hydrochloride solution in 0.1 M sodium cacodylate buffer or phosphate buffer. Protect this solution from light and cool it down on ice to 4 ^oC in order to increase the solubility of oxygen. Saturate the solution with oxygen by bubbling oxygen through the solution (use a hose that is inserted into the solution and connected to an oxygen bottle). Check the pH again and adjust if necessary.
   NOTE: It is crucial for the photooxidation reaction that the pH is between pH 7.0 and pH 7.6.
7. Mount the dish with the fixed cells carefully onto an inverted fluorescence microscope equipped with a 40x oil immersion objective lens and move it to the area of interest. Excite the miniSOG tag with blue light by using filter cubes for GFP or CFP. MiniSOG has an excitation maximum at 448 nm with a shoulder at 473 nm¹⁰.
8. Continue with the illumination even after the green miniSOG fluorescence has completely disappeared and until the brown photo-oxidation product of the polymerized diaminobenzidine emerges in the transmitted light channel. When the oxidation product is visible in most of the cells, turn off the fluorescence illumination to stop the photo-oxidation.
   NOTE: Photo-oxidation can take up to several minutes depending on the objective lens, filter transmission wavelengths and efficiency, and the illumination source.
9. Postfix the cells with 1 ml of 2% glutaraldehyde in 0.1 M sodium cacodylate pH 7.4 buffer or phosphate buffer pH 7.4 for 30 min.
10. Fix the membranes of the cells with 0.1%-0.5% osmium tetroxide for 20 min in 0.1 M sodium cacodylate buffer pH 7.4 or in 0.1 M phosphate buffer pH 7.4.
    NOTE: It is recommended to use the lowest possible concentration of osmium tetroxide required to stabilize the membranes. Since the polymerized DAB precipitates have a high density of nitrogen, which can be detected directly by ESI, a strong osmium staining is not necessary to identify the miniSOG-tagged protein and might be detrimental to the ESI. Osmium tetroxide is very toxic! Work in a fume hood and dispose the toxic waste adequately.
11. Dehydrate, the cells through an ethanol series using 30%, 50%, 70%, 90%, 98% 100% ethanol steps (each 5 min). Subsequently, incubate the cells in a 1:1 mix of 100% ethanol and acrylic resin (LR White) and put the dish onto a shaker for 4 hr to facilitate infiltration into the cells before incubating the cells for at least another 4 hr in 100% acrylic resin (LR White).
12. In order to polymerize the resin, cut off the lid of a labeled 2 ml microcentrifuge tube with a razor blade and coat the rim with acrylic resin accelerator. Make sure that the accelerator only covers the rim and does not flow into the tube. Subsequently, carefully fill the tube approximately two-thirds with LR White resin using a Pasteur pipette. Take care that the resin does not get into contact with the accelerator on the rim!
13. Remove the resin that infiltrated the cells in the dish and place the dish upside down onto the upright standing microcentrifuge tube so that the width of the tube almost completely fills up the glass covered observation window of the dish.
14. Wait 1 to 2 min for the accelerator to seal the tube and the coverslip, then invert the tube so that the resin in the tube now covers the cells.
15. Place the dish with the tube into an oven at 60 °C and cure it for 12 hr.
16. When the block is cured, remove the dish with the attached resin-filled microcentrifuge tube from the oven and separate the microcentrifuge tube from the dish. Facilitate separation by freeze-thaw cycles in liquid nitrogen and hot water.
17. Discard the glass bottom dish and carefully cut open the microcentrifuge tube with a razor blade in order to remove the block.
18. Label the block with a permanent marker.
19. Use a razor blade to trim the block so that nothing but the 1 mm² region that contains the area previously marked with the diamond or tungsten pen (or contains the area with the cells of interest) remains.
20. Mount the block in an ultramicrotome, trim the block with a trimming knife and cut ultra-thin sections of approximately 50 nm using a diamond knife.
21. Pick up the sections on high-transmission 300 mesh grids. These grids have very small grid bars and thus cover fewer cells in the area of interest.
22. Coat the sections on the grids with about 0.2-0.4 nm of carbon using a carbon coater to help stabilize them under the electron beam of the TEM.

5. Electron Microscopy

1. Load the grids into a TEM that is equipped with an energy filter. After tuning the microscope and the energy filter according to the manufacturer’s instructions, inspect the cells on the sections in the low magnification mode (normal transmission) and compare them to fluorescence data when appropriate (obtained in step 4.4).
2. Once a nucleus with interesting features (DNA damage track or DNA repair foci) is found, switch to the energy filtering mode and record a thickness map.
   NOTE: This procedure includes the recording of a zero-loss and an unfiltered image. Thicknesses up to 0.3 mean free path (30% of the electrons of the incident beam gets scattered by the sample) are thin enough to generate good elemental maps.
3. Record ratio maps of the elements phosphorus and nitrogen using the software that controls the energy filter of the microscope used. Record the phosphorus map post edge images at 175 eV energy loss with a slit width of 20 eV and pre-edge images at 120 eV energy loss, also with a slit width of 20 eV. For the nitrogen maps, record post edge images at 447 eV with a slit width of 35 eV and pre-edge images at 358 eV, also with a slit width of 35 eV.
   NOTE: Since the content of the imaged elements (phosphorus and nitrogen) is relatively low (approx. 1%), choose “Ratio Image” (qualitative elemental map) over the “3 window method” (quantitate elemental map) in order to generate images with a better signal to noise ratio.

6. Image Processing

1. Open the elemental ratio maps in an image processing software that can handle the file format of the acquired pictures (e.g., Digital Micrograph). Copy the nitrogen map into the red channel and the phosphorus map into the green channel of an RGB image, then superimpose and align the maps.
2. Export the composite image as a tagged image file format (TIFF) file. Open the image in an image processing software that can use layers (e.g., Photoshop).
3. Adjust the dynamic range of each channel by rescaling the minimum and maximum values in the image to an 8-bit data set (0 to 255).
4. Subtract the phosphorus content from the nitrogen map (using the “Layers” window) in order to generate a qualitative map that shows the distribution of protein.
5. Convert the maps of the nucleic acid (phosphorus) and the protein (nitrogen) created in the previous step into “indexed color” and create a yellow lookup table in the CMYK color space for the map showing the nucleic acid distribution and a cyan lookup table to show the non-nucleoprotein map. The colors are chosen to generate maximum contrast between the two elemental maps.
6. Create a new image and import the maps showing the distributions of phosphorus and protein as different layers. Use the “screen” transparency mode in order to see both layers superimposed on each other.
7. Open the zero-loss image acquired in step 5.2. This image represents an image of the recorded area that looks like a conventional TEM image but only contains the electrons that have passed through the specimen without colliding with the specimen. Export this image as a TIFF image.
8. Open the exported zero loss image in a photo editor and copy it into the uppermost layer of the image showing the elemental map. Align the image using the “screen” transparency mode.
9. Optionally threshold the zero-loss image to segment the signal that originates from the miniSOG. Add the resulting image as a separate layer as described above.

Wyniki

ESI

Comparing the ESI images of the nucleus (Figure 1) with the conventional TEM images (e.g., Figure 7-1) reveals a dramatic increase in anatomical structures that can be easily distinguished. The chromatin ridges appear in yellow and it is quite easy to identify the chromocenters in mouse cells. Nucleoli can easily be distinguished from chromatin by their round structure and different color, since the pre-assembled ribosomes al...

Dyskusje

ESI can serve as an excellent tool for examining different states of chromatin and ribonucleoproteins in the nucleus because it is able to specifically map areas that are rich in phosphorus. It profoundly augments the amount of detail that can be obtained by an electron microscope and is not dependent on nonspecific contrasting methods. One disadvantage of ESI is that it requires thinner sections than conventional TEM at the same accelerating voltage. This can be overcome by working with serial sections or using tomograp...

Materiały

Name	Company	Catalog Number	Comments
35 mm glass bottom dishes	MatTek	P35G-1.5-14-C
Gridded 35 mm glass bottom dishes	MatTek	P35G-2-14-CGRD	not made for immersion objectives
LR White: acryl resin	emsdiasum	14381
LR White accelerator	emsdiasum	14385
3,3′-Diaminobenzidine tetrahydrochloride 10 mg tablets	Sigma	D5905
Slim Bar Grids 300 Mesh	SPI	1161123
Tungsten-Point Lab Pen	emsdiasum	41148
Osmium Tetroxide	emsdiasum	19100
Carbon Coater 208 carbon	Cressington
Ultra microtome Leica EM UC6	Leica
Photoshop CS5	Adobe		might even work with older versions
Digital Micrograph V.2.30	Gatan		might even work with older versions
Hoechst 33342	Sigma	H-1399
Effectene	Quiagen
MiniSOG-Constructs	Tsien-Lab		tsienlab@yahoo.com
MDC1 miniSOG mCherry
53BP1 miniSOG mCherry
Rad52 miniSOG mCherry
cesium 137 radiation source "MARK 1"	(J.L. Shepherd & Associated)
ImageJ/FiJi	open source		http://fiji.sc/Fiji
2 ml Eppendorf tubes	Fisherbrand	05-408-146
Diamond Knife ultra 35°	Diatome
Trimming Knife ultratrim	Diatome
Sodium cacodylate trihydrate	emsdiasum	12300	Caution Toxic!
Glutaraldehyde EM Grade 8%	emsdiasum	16020	Caution Toxic!
Sodium phosphate dibasic	emsdiasum	21180
Sodium phosphate monobasic	emsdiasum	21190
Paraformaldehyde	emsdiasum	19202
Osmium tetroxide 4% solution	emsdiasum	19150
Inverted Fluorescence Micoscope Axiovert 200M	Zeiss
Hydrochloric Acid	Fisherbrand	A142-212
Sodium Hydroxide Solution 10M	Fluka	72068
Oxygen	Medigas
3-Amino-1,2,4-triazole	Sigma	A8056
Potassium cyanide	Sigma	207810	Caution Toxic!
Ethanol	emsdiasum	15058	Caution Toxic!
Razor blade Single Edge Carbon Steel	emsdiasum	71960	Caution Sharp!
DMEM	Sigma	D 5546
FBS	Life Technologies	16000-044
G418	Life Technologies	11811-023
DMSO	Sigma	D2650
Transmission Electron Microscope 200 kV	JEOL	2100
GIF Tridiem 863 Energy filter	Gatan