The authors thank Tammo Reisewitz for critical reading of the manuscript, and both Mako Hayashi and Sayuri Hagiwara for careful cultivation and observation of cyanobacteria, Yoshitaka Kimori for image processing, Chihong Song, Naoyuki Miyazaki, and Miyoko Nagayoshi for helping with segmentation of the structures. This work was supported by the Collaborative Study Program of the National Institute for Physiological Sciences (NIPS) to Y.K.

1. Synchronous Culture of Cyanobacteria
<ol>
	<li>Culture S. elongatus PCC 7942 on sterilized BG 11 plate (in a 9 cm sterile plastic petri dish) containing 1.5% (w/v) agar and 0.3% (w/v) sodium thiosulfate8.</li>
	<li>Place the plates in a growth chamber at 23 &#176;C with a light intensity of 50 &#181;E/m2/s and subject to 12 h light/12 h dark cycles.</li>
	<li>Transfer the cells onto fresh BG11 agar plates once a week. 
	NOTE: The cultures on the agar will appear as green bands of actively proliferating cells after one week under this culture condition.</li>
	<li>Take green clumps of cells with a flame sterilized wire loop and streak the cells onto a fresh BG11 agar plate. Do this on a clean bench.</li>
</ol>
2. Monitoring by Fluorescence Microscopy
<ol>
	<li>Use cells cultured on the agar plate for 6 days to observe DNA compaction. Collect cells from the plate at the end of the light period by pouring 1 mL of 0.2 M sucrose solution over the cells. Repeat pouring the solution onto the cells so that most of the cells are collected. Transfer the suspended-cell solution into a micro tube for DNA staining.</li>
	<li>Add DNA staining dye (e.g. Hoechst 33342) solution to 500 &#181;L of the suspended cell solution in a micro tube to a final concentration of 1 &#181;g/mL. Then, keep the tube in the dark for 10 min.</li>
	<li>Centrifuge for 1 min at 2,000 x g to sediment the cells. Discard the supernatant and add 10 &#181;L of 0.2 M sucrose solution to obtain a dense cell suspension.</li>
	<li>Transfer 1 &#181;L of the solution containing stained cells to a slide glass, put a cover slip and observe with a fluorescence microscope equipped with a UV filter using an objective lens with a magnification of 100X and immersion oil.</li>
	<li>Confirm that the DNA compaction is observed at this point in most cells, then prepare the sample for the next freezing step.</li>
</ol>
3. Sample Freezing for Cryo-HVET
<ol>
	<li>Set up a plunge-freezing device. Fill the tank with liquid nitrogen and start cooling the cryo-chamber after connecting the tank and the chamber with a Teflon tube.</li>
	<li>Fill liquid ethane into a small cupper pot inside the chamber after cooling down the chamber to liquid nitrogen temperature. Wear glasses during operation, because liquid ethane is explosive.</li>
	<li>Glow discharge the carbon-side of a holey carbon-coated EM grid (holey grid) for 30 s at 50 mA using a plasma ion bombarder.</li>
	<li>Apply 1 &#181;L of BSA gold tracer (15 nm) to the holey grid as a fiducial marker.</li>
	<li>Apply a 2.5 &#181;L aliquot of cells in the DNA compaction stage to the holey grid. Blot off the excess solution with a filter paper. Plunge the grid into liquid ethane using a plunger in the plunge freezing device immediately.</li>
	<li>Store the frozen grids in liquid nitrogen storage until they are examined.</li>
</ol>
4. Cryo-HVET
<ol>
	<li>Set up the HVEM at a high voltage of 1 MV.</li>
	<li>Mount the frozen grid in a cryo-specimen holder for HVEM precooled to -150 &#176;C with liquid nitrogen inside the cryo-workstation, and load it into the HVEM. Be careful to avoid contamination by ice.</li>
	<li>Select an imaging area at a lower magnification of 1,000X. Adjust the eucentric z-axis height.</li>
	<li>Tilt the specimen stage to -60&#176; and remove the backlash of the tilting rotation.</li>
	<li>Adjust the focus near the target location at a magnification of 10,000X. Set an under focus of 6 to 10 &#181;m by deviation from the focused image. For imaging, set the dose to 2 e-/&#197;-2 or less on the specimen beforehand.</li>
	<li>Measure the electron dose by the current density on the image screen in HVEM. Take an image on an electron film or by digital camera at the same magnification as in the focusing process.</li>
	<li>Collect tilt images manually by the same procedure as in (4.5) from -60&#176; to +60&#176; at a tilt angle increment of 2&#176; to 4&#176;. 
	NOTE: In many modern electron microscopes, the acquisition of the tilt series is automated by the combination of a digital camera. In that case, follow the instruction manual. For negative films, develop the films for 12 min at 20&#176;C in a developer tank using a full-strength developer and fix in a fixer for 10 min. Digitize the films at a resolution of 4,000 dpi (0.635 nm/pixel on the image) using a flatbed scanner. In the case of a digital camera, subject the collected images directly to the next image processing step. If necessary, reduce the image size by a median filter using two to four binning (size reducing factor) and appropriate software (e.g. ImageJ).</li>
</ol>
5. Tomographic Reconstruction
<ol>
	<li>Make a stack image file from the individual tilt images using the command &#34;tif2mrc&#34; or &#34;newstack&#34; in IMOD software9.</li>
	<li>Start eTomo GUI software in IMOD and set image parameters: pixel size, fiducial diameter, image rotation, etc. Then create scripts.</li>
	<li>Execute individual programs according to the software listed in tables of materials, where the tilt series is aligned using fiducial markers (with a mean residual error less than 0.5). Finally, reconstruct a 3D tomogram using the SIRT algorithm in IMOD.</li>
	<li>Extract a region of interest (ROI) from the tomogram and denoise using a denoise filter: an anisotropic diffusion filter in IMOD, a bilateral filter in EMAN10, or a mathematical morphology filter11, etc., with appropriate parameters to enhance the contrast.</li>
</ol>
6. Segmentation of the Feature of Interest
NOTE: The procedure described below is specific to the software used (see Materials Table) but other software packages can be used instead. Refer to their user guide.
<ol>
	<li>In the 3D Viewer window, open the tomogram file on Amira software and generate an OrthoSlice.</li>
	<li>In the Segmentation Editor window, create a segmentation file by selecting a new &#34;label field&#34;.</li>
	<li>Manually trace the border of the feature of interest (FOI). Follow the FOI through all tomogram slices. For the second FOI, create a new &#34;material&#34; and repeat the same operation.</li>
	<li>Generate a surface rendering by selecting the &#34;SurfaceGen&#34; menu. To visualize the segmented volume, select the &#34;SurfaceView&#34; menu. To move, rotate, and zoom in the 3D volume, use the tools in the 3D Viewer window.</li>
	<li>For automatic segmentation, use the Magic Wand Tool. Click on an object, and adjust the sliders in Display and Masking to cover the range of values so that the object is fully selected by its features.</li>
</ol>

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The authors declare no competing financial interests.

We have presented a sequence of protocols for visualizing transient DNA compaction in cyanobacteria. The basic concept is similar to that of correlative light and electron microscopy (CLEM)12. In addition, in this method, live cyanobacteria were monitored by fluorescence microscopy, rapidly frozen on EM grids, and directly visualized with high voltage cryo-electron tomography. As a first application, the detailed structure of DNA compacted bacterial cells was successfully visualized in 3D. Currently, this procedure is specific to this subject, but it will be applied more extensively, with modified methodology in some cases. Here, the advantages, the limitations, and the future possibilities of this method are discussed.
One of the advantages of this method is the 3D visualization of the whole cell. 1 MV HVEM successfully visualized the dynamic structure of the subcellular organelles in the DNA compacted cells. However, the fine structure inside normal cells could not be distinguished due to low image contrast. Increasing inelastic and multiple scattering in thick specimens blurs the image13. Zero-loss and most-probable-loss image filtering by an energy filter can improve image contrast by reducing inelastic scattering14,15, but it will not work for specimens thicker than iMFP. The zero-loss and most-probable-loss peaks drastically decrease with specimen thickness. It is particularly difficult to obtain a sufficient signal-to-noise ratio for the electron sensitive ice-embedded specimens. Murata et al. have shown that 1MV scanning transmission microscopy (STEM) gives higher image contrast than a bright field image in plastic embedded yeast cells with 5 &#181;m thickness, where the image contrast is primarily given by amplitude contrast13. However, it is expected that the effect of knock-on damage on higher accelerated electrons creates another limitation to the irradiation dose for damage-sensitive cryo-specimens16. The application of Volta and Zernike phase plates17,18 for HVEM may be able to reduce knock-on damage by reducing the total dose in the future. Another limitation to using HVEM for thick specimens comes from the fact that the user facilities that provide HVEM are scarce worldwide.
Using an alternative methodology to observe thick specimens, cryo-STEM tomography at 300 kV has demonstrated high contrast images on frozen-hydrated specimens with thickness in excess of several hundred nanometers19. To retrieve phase contrast in cryo-STEM, electron pthychographic microscopy has also been introduced, in which the phase plate in the condenser lens transposes a phase-modulated diffraction to a pixelated 2D detector20. The images are retrieved by calculation from multiple diffractions. For direct and fast 3D cryo-imaging of large native frozen samples, cryo-FIB-SEM can also be used21, where serial sectioning with a focused ion beam and block face imaging is applied for imaging fully hydrated frozen specimens. Although these technologies expand the viewing range of biological specimens, it is difficult to find the target location of the bacteria, e.g. labeled bacteria, because the target is completely under the ice and cannot be identified before trimming.
DNA compaction produces a distinct structure in cyanobacteria. DNA compacted cells are readily distinguished even without being stained due to a large density bias within the cells that is not present in normal cells. However, in order to visualize more local events within the cell, it is necessary to transfer the fluorescently labeled ROI into the electron microscope. For correlative light and electron microscopy (CLEM), light microscope images and electron microscope images are generally correlated using fluorescent latex beads or quantum dots on EM finder grids12. The labelling particles must be of high electron density in addition to fluorescence. They can accurately and reliably correlate the positions between the two images. Furthermore, by confirming the labeled area with cryo-light microscopy, complete overlap of ROI can be achieved between the two microscopes. When characterizing more detailed structural events in DNA compaction, these particles and cryo-light microscope will be an indispensable tool for a more robust and accurate correlation in the future.
This article shows how to characterize the transient structure of DNA compaction in cyanobacteria by a combination of synchronous culture, fluorescence microscopy and high voltage cryo-electron tomography. This protocol focuses on the observation of compacted DNA. By combining this method with other new technologies mentioned above, it will be possible to investigate the process of DNA compaction in further detail, and suitably modified methods are widely applicable to other dynamic structural events in bacteria.

DNA compaction is a dramatic cytoplasmic event that has been identified in some cyanobacteria. When Synechococcus elongatus was cultured under 12 h each light/dark cycle, DNA appeared condensed at the end of the light period, which was clearly different from its appearance at the other time points1. It has been suggested that this process is controlled by a circadian clock based on the Kai proteins2. Seki et al. have reported that the DNA stained with Hoechst 33342 was compacted in S. elongatus cells toward the end of the light period and showed a wavy rod-shape under a fluorescence microscope. The compacted DNA then separated into two at the center of the rod as the cell divided, and finally returned to a normal uniform distribution in each daughter cell3. However, its transient nature and large size for electron microscopy impeded structural analysis. Murata et al. combined several methods, including synchronous culture, fluorescence microscopy, rapid freezing, and high voltage electron cryo-tomography (cryo-HVET), and succeeded in identifying the structure of transient DNA compaction, including the kinetics of polyphosphate bodies (PPBs)4. The manuscript provides visual explanation of such a difficult material in detail by combining the experimental procedures.
S. elongatus has a capsule shape, a length of 2 to 5 &#181;m, a width of about 0.5 &#181;m, and the perfect DNA compaction appears in living cells only for a very short time. Therefore, the structural changes occurring in the cyanobacterial DNA compaction were unknown in detail. In order to investigate these structures by electron microscopy, it is necessary to overcome two main technical problems. One is the observation of such a thick specimen of the whole bacterium at near native conditions, and the other is the rapid fixation of a dynamic structure. As for the first problem, the inelastic mean free path (iMFP) of electrons depends on the accelerating voltage of the electron microscope5. In a transmission electron microscope (TEM) of 300 kV, it is less than 350 nm. For example, when an ice-embedded cyanobacterium (specimen thickness &#8776; 600 nm) is observed in 200kV TEM (iMFP &#8776; 250 nm), the structures inside cell are difficult to observe. By contrast, 1 MV TEM (iMFP &#8776; 500 nm) can give and image of the cytoplasmic structure throughout the cell (Figure 1). In this protocol, as one part of the solution, a high voltage electron microscope (HVEM) at an accelerating voltage of 1 MV was employed. However, facilities that implement HVEM are limited worldwide. Possible alternative solutions are also discussed in the Discussion section. The second problem was solved by cryo-electron microscopy (cryo-EM). This is a powerful tool for visualizing dynamic structures at near native conditions, where the specimen is rapidly frozen in liquid ethane using a rapid freezing device, and the frozen moment is directly observed with a minimum of modifications6. Combining with tomography, a snapshot of three-dimensional (3D) structures can be reconstructed from the tilt series7. In this experiment, DNA compaction was reproduced in S. elongatus using synchronous culture under 12 h each light/dark cycle, and the timing of the freezing of the specimen was determined by monitoring under a fluorescence microscope.
The approaches described here are widely applicable to the study of dynamic structures in bacterial cells, e.g. cell division, chromosome segregation and phage infection, and have a potential to open new avenues in microbiological research.

<table border="0" cellpadding="0" cellspacing="0" width="796">
	<tbody>
		<tr height="26">
			<td height="26" style="height:26px;width:235px;">Hoechst 33342 solution</td>
			<td style="width:143px;">Dojindo</td>
			<td style="width:155px;">346-07951</td>
			<td style="width:263px;">1mg/mL in H2O</td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">Agar Powder (for plant culture)</td>
			<td>Wako</td>
			<td>016-11875</td>
			<td></td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">Boric acid</td>
			<td>Wako</td>
			<td>027-02192</td>
			<td>99.5%</td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">Manganese chloride tetrahydrate</td>
			<td>Wako</td>
			<td>139-00722</td>
			<td>99%</td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">Zinc sulfate heptahydrate</td>
			<td>Wako</td>
			<td>264-00405</td>
			<td>99.5%</td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">Sodium molybdate</td>
			<td>Wako</td>
			<td>196-02472</td>
			<td>99%</td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">Copper sulfate pentahydrate</td>
			<td>Wako</td>
			<td>039-04412</td>
			<td>99.5%</td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">Cobalt nitrate hexahydrate</td>
			<td>Wako</td>
			<td>031-03752</td>
			<td>99.5%</td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">Sodium nitrate</td>
			<td>Wako</td>
			<td>191-02542</td>
			<td>99%</td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">Magnesium sulfate heptahydrate</td>
			<td>Wako</td>
			<td>131-00405</td>
			<td>99.5%</td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">Calcium chloride dehydrate</td>
			<td>Wako</td>
			<td>031-00435</td>
			<td>99.5%</td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">Citric acid</td>
			<td>Wako</td>
			<td>036-05522</td>
			<td>98%</td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">EDTA-2Na</td>
			<td>Dojindo</td>
			<td>343-01861</td>
			<td>99.5%</td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">Sodium carbonate</td>
			<td>Wako</td>
			<td>197-01581</td>
			<td>99.8%</td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">Potassium phosphate dibasic</td>
			<td>Wako</td>
			<td>164-04295</td>
			<td>99%</td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">TES (Good&#8217;s buffer)</td>
			<td>Dojindo</td>
			<td>344-02653</td>
			<td>99%</td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">Ferric ammonium citrate</td>
			<td>Wako</td>
			<td>092-00802</td>
			<td>1st Grade</td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">Sodium thiosulfate pentahydrate</td>
			<td>Wako</td>
			<td>197-03585</td>
			<td>99%</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">BSA gold tracer 15nm</td>
			<td>Aurion</td>
			<td>215.133</td>
			<td></td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">Quantifoil EM grid</td>
			<td>Quantifoil MicroTools</td>
			<td>R3.5/1 Copper grid</td>
			<td></td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">Electron films</td>
			<td>Kodak</td>
			<td>SO-163</td>
			<td></td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">Developer</td>
			<td>Kodak</td>
			<td>D19</td>
			<td></td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">Fixer</td>
			<td>Kodak</td>
			<td>Rapid fixer</td>
			<td>Solution</td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">Filter paper</td>
			<td>Whatman</td>
			<td>Grade 1</td>
			<td></td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">Growth chamber</td>
			<td>NKsystem</td>
			<td>LH-100SP</td>
			<td></td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">Fluorescent microscope</td>
			<td>Nikon</td>
			<td>ECLIPSE 50i</td>
			<td></td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">High voltage TEM</td>
			<td>HItachi</td>
			<td>H1250M</td>
			<td></td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">Cryo-specimen holder for HVEM</td>
			<td>Gatan</td>
			<td></td>
			<td></td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">plunge-freezing device</td>
			<td>Leica</td>
			<td>EM CPC</td>
			<td></td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">Plasma Ion bombarder</td>
			<td>Vacuum device</td>
			<td>PIB-10</td>
			<td></td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">Liquid nitrogen storage</td>
			<td>Taylor-Wharton</td>
			<td>25LDB</td>
			<td></td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">Developing tank</td>
			<td>Dosaka EM</td>
			<td>TB-3-75</td>
			<td></td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">flatbed scanner</td>
			<td>Nikon</td>
			<td>Coolscan 9000ED</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Segmentation software</td>
			<td>FEI</td>
			<td>Amira</td>
			<td>https://www.fei.com/software/amira</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Tomographic Reconstruction software</td>
			<td>eTOMO</td>
			<td></td>
			<td>http://bio3d.colorado.edu/imod</td>
		</tr>
	</tbody>
</table>

visualization of dna compaction in cyanobacteria by high-voltage cryo-electron tomography

This protocol describes how to visualize the transient DNA compaction in cyanobacteria. DNA compaction is a dramatic cytoplasmic event recently found to occur in some cyanobacteria before cell division. However, due to the large cell size and the transient character, it is difficult to investigate the structure in detail. To overcome the difficulties, first, DNA compaction is reproducibly produced in the cyanobacterium Synechococcus elongatus PCC 7942 by synchronous culture using 12 h each light/dark cycle. Second, DNA compaction is monitored by fluorescence microscopy and captured by rapid freezing. Third, the detailed structure of DNA compacted cells is visualized in three dimensions (3D) by high voltage cryo-electron tomography. This set of methods is widely applicable to investigate transient structures in bacteria, e.g. cell division, chromosome segregation, phage infection etc., which are monitored by fluorescence microscopy and directly visualized by cryo-electron tomography at appropriate time points.

In a precise synchronous culture under 12 h each light/dark cycle, DNA labeled with Hoechst shows a normal uniform distribution in the dark condition (Figure 2a). However, it progressively compacts within the cell during the light period, and appears as a wavy rod-like structure (Figure 2b) at the end of the light period. Finally, the rod divides at the center (arrows in Figure 2c) and its two parts are distributed into the daughter cells (Figure 2d). After cell division, the compacted DNA disappears immediately, and the DNA returns to a normal uniform distribution.
When an aliquot of cells containing the cells in the final stage of DNA compaction were immediately transferred onto a holey grid and rapid frozen in liquid ethane, and the frozen grid was observed by 1 MV cryo-HVEM, the internal structures of cyanobacteria including DNA, thylakoid membrane layers, cell walls, and PPBs, appeared as in a snapshot at the moment of freezing (Figure 3a). Many cells showed distinct DNA compaction in the cells (white arrows in Figure 3a), and could be easily distinguished from normal cells (white arrowheads in Figure 3b). Some exhibited a constriction at the center of the cells as expected before cell division (yellow arrow in Figure 3a).
In 3D tomograms, major organelles of the cell could be segmented; cell wall, thylakoid membrane layers, DNA, and PPBs could be distinguished (Figure 4). In particular, the compacted DNA was separated by a distinct gap in the cytoplasm where the DNA was surrounded by low density material and the thylakoid membrane layers were distorted along the wavy rod of compacted DNA. Dynamic behavior of PPBs was newly observed: in DNA compacted cells, many small PPBs were seen to adhere to DNA, whereas they are large and fewer in normal cells. In addition, most of the PPBs appeared as pairs, and some DNA appeared to be in a process of separation from the PPBs. This suggested that PPBs themselves are divided into two by DNA duplication and function as suppliers of phosphate for DNA synthesis.
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/57197/57197fig1.jpg" /> 
Figure 1. Cryo-EM images of ice-embedded normal cyanobacteria, S. elongatus PCC 7942 at different accelerating voltages. a) 200kV and b) 1000kV. Scale bar = 500 nm. <a href="https://cloudflare-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/57197/57197fig1large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/57197/57197fig2.jpg" /> 
Figure 2: Fluorescence microscopy of cyanobacterium S. elongatus cells. After synchronous culture under 12 hour each light/dark cycles, cells were stained with Hoechst 33342. (a) Cells after 2 h of onset of the dark state show uniform DNA labeling. The DNA in many cells gradually condensed during the light period. It ultimately formed a thick wavy rod (b) typical of DNA compaction. After this stage, the compacted DNA structures quickly divided at their centers (arrowheads) during cell division (c), and the two fragments separated into the daughter cells (d). The daughter cells returned to uniform DNA labeling again in the dark period. Scale bar = 2 &#181;m. <a href="https://cloudflare-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/57197/57197fig2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 3" class="xfigimg" src="/files/ftp_upload/57197/57197fig3.jpg" /> 
Figure 3: Cryo-HVEM image of cyanobacteria embedded in amorphous ice. (a) shows a raw image at 0&#176; tilt. Images were taken at the end of the light period. Some cells show wavy rod-shaped DNA bodies similar to eukaryotic condensed chromosomes (white arrows). In normal cells, cyanobacteria exhibit a discernable DNA structure within the cytoplasm (white arrowheads). Some exhibit a constriction at the center of the cells as expected before cell division (yellow arrow). (b) xy, xz, yz-slices of a 3D tomogram. Yellow dotted lines show intersections of slices. Scale bar = 2 &#181;m. <a href="https://cloudflare-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/57197/57197fig3large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 4" class="xfigimg" src="/files/ftp_upload/57197/57197fig4.jpg" /> 
Figure 4: Ultrastructure of cells containing compacted DNA. The major components are segmented: cell wall (bright yellow), thylakoid membranes (red), and DNA (blue). (a) shows a z-slice of a 3D tomogram. (b) all segments. The constriction indicating cell separation appears in the center of the cell (arrow). PPBs are modeled as yellow or orange spheres; each orange sphere represents the counterpart of the closest yellow sphere. Scale bar = 500 nm. <a href="https://cloudflare-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/57197/57197fig4large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

Watch this Scientific Journal Video about Visualization of DNA Compaction in Cyanobacteria by High-voltage Cryo-electron Tomography at JoVE.com

Visualization of DNA Compaction in Cyanobacteria by High-voltage Cryo-electron Tomography

This protocol describes how to visualize the transient DNA compaction in cyanobacteria. Synchronous cultivation, monitoring by fluorescence microscopy, rapid freezing, and high voltage cryo-electron tomography are used. A protocol for these methodologies is presented, and future applications and developments are discussed.

This protocol describes how to visualize the transient DNA compaction in cyanobacteria. DNA compaction is a dramatic cytoplasmic event recently found to ...

visualization-dna-compaction-cyanobacteria-high-voltage-cryo-electron

Nanyang Technological University

Research

JoVE Journal

Biology

9.0K Views.  National Institute for Physiological Sciences. This protocol describes how to visualize the transient DNA compaction in cyanobacteria. Synchronous cultivation, monitoring by fluorescence microscopy, rapid freezing, and high voltage cryo-electron tomography are used. A protocol for these methodologies is presented, and future applications and developments are discussed.

Video: Visualization of DNA Compaction in Cyanobacteria by High-voltage Cryo-electron Tomography

Visualization of DNA Replication in the Vertebrate Model System DT40 using the DNA Fiber Technique

Maintenance of replication fork stability is of utmost importance for dividing cells to preserve viability and prevent disease. The processes involved not only ensure faithful genome duplication in the face of endogenous and exogenous DNA damage but also prevent genomic instability, a recognized causative factor in tumor development.
Here, we describe a simple and cost-effective fluorescence microscopy-based method to visualize DNA replication in the avian B-cell line DT40. This cell line provides a powerful tool to investigate protein function in vivo by reverse genetics in vertebrate cells1. DNA fiber fluorography in DT40 cells lacking a specific gene allows one to elucidate the function of this gene product in DNA replication and genome stability. Traditional methods to analyze replication fork dynamics in vertebrate cells rely on measuring the overall rate of DNA synthesis in a population of pulse-labeled cells. This is a quantitative approach and does not allow for qualitative analysis of parameters that influence DNA synthesis. In contrast, the rate of movement of active forks can be followed directly when using the DNA fiber technique2-4. In this approach, nascent DNA is labeled in vivo by incorporation of halogenated nucleotides (Fig 1A). Subsequently, individual fibers are stretched onto a microscope slide, and the labeled DNA replication tracts are stained with specific antibodies and visualized by fluorescence microscopy (Fig 1B). Initiation of replication as well as fork directionality is determined by the consecutive use of two differently modified analogues. Furthermore, the dual-labeling approach allows for quantitative analysis of parameters that influence DNA synthesis during the S-phase, i.e. replication structures such as ongoing and stalled forks, replication origin density as well as fork terminations. Finally, the experimental procedure can be accomplished within a day, and requires only general laboratory equipment and a fluorescence microscope.

DT40, a model vertebrate genetic system, provides a powerful tool to analyze protein function. Here we describe a simple method that allows qualitative analysis of parameters that influence DNA synthesis during the S-phase in DT40 cells at the single molecule level.

Maintenance of replication fork stability is of utmost importance for dividing cells to preserve viability and prevent disease. The processes involved not ...

Visualization of ATP Synthase Dimers in Mitochondria by Electron Cryo-tomography

Electron cryo-tomography is a powerful tool in structural biology, capable of visualizing the three-dimensional structure of biological samples, such as cells, organelles, membrane vesicles, or viruses at molecular detail. To achieve this, the aqueous sample is rapidly vitrified in liquid ethane, which preserves it in a close-to-native, frozen-hydrated state. In the electron microscope, tilt series are recorded at liquid nitrogen temperature, from which 3D tomograms are reconstructed. The signal-to-noise ratio of the tomographic volume is inherently low. Recognizable, recurring features are enhanced by subtomogram averaging, by which individual subvolumes are cut out, aligned and averaged to reduce noise. In this way, 3D maps with a resolution of 2 nm or better can be obtained. A fit of available high-resolution structures to the 3D volume then produces atomic models of protein complexes in their native environment. Here we show how we use electron cryo-tomography to study the in situ organization of large membrane protein complexes in mitochondria. We find that ATP synthases are organized in rows of dimers along highly curved apices of the inner membrane cristae, whereas complex I is randomly distributed in the membrane regions on either side of the rows. By subtomogram averaging we obtained a structure of the mitochondrial ATP synthase dimer within the cristae membrane.

We present a protocol of how to collect and process electron cryo-tomograms of whole mitochondria. The technique provides detailed insights into the structure, function, and organization of large membrane protein complexes in native biological membranes.

Electron cryo-tomography is a powerful tool in structural biology, capable of visualizing the three-dimensional structure of biological samples, such as ...

Cryo-electron Microscopy Specimen Preparation By Means Of a Focused Ion Beam

Here we present a protocol used to prepare cryo-TEM samples of Aspergillus niger spores, but which can easily be adapted for any number of microorganisms or solutions. We make use of a custom built cryo-transfer station and a modified cryo-SEM preparation chamber2. The spores are taken from a culture, plunge-frozen in a liquid nitrogen slush and observed in the cryo-SEM to select a region of interest. A thin lamella is then extracted using the FIB, attached to a TEM grid and subsequently thinned to electron transparency. The grid is transferred to a cryo-TEM holder and into a TEM for high resolution studies. Thanks to the introduction of a cooled nanomanipulator tip and a cryo-transfer station, this protocol is a straightforward adaptation to cryogenic temperature of the routinely used FIB preparation of TEM samples. As such it has the advantages of requiring a small amount of modifications to existing instruments, setups and procedures; it is easy to implement; it has a broad range of applications, in principle the same as for cryo-TEM sample preparation. One limitation is that it requires skillful handling of the specimens at critical steps to avoid or minimize contaminations.

Cryo Electron Microscopes, either Scanning (SEM) or Transmission (TEM), are widely used for characterization of biological samples or other materials with a high water content1. A SEM/Focused Ion Beam (FIB) is used to identify features of interest in samples and extract a thin, electron-transparent lamella for transfer to a cryo-TEM.

Here we present a protocol used to prepare cryo-TEM samples of Aspergillus niger spores, but which can easily be adapted for any number of microorganisms ...

Real-time Imaging of Single Engineered RNA Transcripts in Living Cells Using Ratiometric Bimolecular Beacons

The growing realization that both the temporal and spatial regulation of gene expression can have important consequences on cell function has led to the development of diverse techniques to visualize individual RNA transcripts in single living cells. One promising technique that has recently been described utilizes an oligonucleotide-based optical probe, ratiometric bimolecular beacon (RBMB), to detect RNA transcripts that were engineered to contain at least four tandem repeats of the RBMB target sequence in the 3&#8217;-untranslated region. RBMBs are specifically designed to emit a bright fluorescent signal upon hybridization to complementary RNA, but otherwise remain quenched. The use of a synthetic probe in this approach allows photostable, red-shifted, and highly emissive organic dyes to be used for imaging. Binding of multiple RBMBs to the engineered RNA transcripts results in discrete fluorescence spots when viewed under a wide-field fluorescent microscope. Consequently, the movement of individual RNA transcripts can be readily visualized in real-time by taking a time series of fluorescent images. Here we describe the preparation and purification of RBMBs, delivery into cells by microporation and live-cell imaging of single RNA transcripts.

Ratiometric bimolecular beacons (RBMBs) can be used to image single engineered RNA transcripts in living cells. Here, we describe the preparation and purification of RBMBs, delivery of RBMBs into cells by microporation and fluorescent imaging of single RNA transcripts in real-time.

The growing realization that both the temporal and spatial regulation of gene expression can have important consequences on cell function has led to the ...

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

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 &#181;m, 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.

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

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.

The limits to optical resolution and the challenge of identifying specific protein populations in transmission electron microscopy have been obstacles in ...

Visualization of Surface-tethered Large DNA Molecules with a Fluorescent Protein DNA Binding Peptide

Large DNA molecules tethered on the functionalized glass surface have been utilized in polymer physics and biochemistry particularly for investigating interactions between DNA and its binding proteins. Here, we report a method that uses fluorescent microscopy for visualizing large DNA molecules tethered on the surface. First, glass coverslips are biotinylated and passivated by coating with biotinylated polyethylene glycol, which specifically binds biotinylated DNA via avidin protein linkers and significantly reduces undesirable binding from non-specific interactions of proteins or DNA molecules on the surface. Second, the DNA molecules are biotinylated by two different methods depending on their terminals. The blunt ended DNA is tagged with biotinylated dUTP at its 3' hydroxyl terminus, by terminal transferase, while the sticky ended DNA is hybridized with biotinylated complimentary oligonucleotides by DNA ligase. Finally, a microfluidic shear flow makes single DNA molecules stretch to their full contour lengths after being stained with fluorescent protein-DNA binding peptide (FP-DBP).

We present an approach for visualizing fluorescent protein DNA binding peptide (FP-DBP)-stained large DNA molecules tethered on the polyethylene glycol (PEG) and avidin-coated glass surface and stretched with microfluidic shear flows.

Large DNA molecules tethered on the functionalized glass surface have been utilized in polymer physics and biochemistry particularly for investigating ...

Genetic Modification of Cyanobacteria by Conjugation Using the CyanoGate Modular Cloning Toolkit

Cyanobacteria are a diverse group of prokaryotic photosynthetic organisms that can be genetically modified for the renewable production of useful industrial commodities. Recent advances in synthetic biology have led to development of several cloning toolkits such as CyanoGate, a standardized modular cloning system for building plasmid vectors for subsequent transformation or conjugal transfer into cyanobacteria. Here we outline a detailed method for assembling a self-replicating vector (e.g., carrying a fluorescent marker expression cassette) and conjugal transfer of the vector into the cyanobacterial strains Synechocystis sp. PCC 6803 or Synechococcus elongatus UTEX 2973. In addition, we outline how to characterize the performance of a genetic part (e.g., a promoter) using a plate reader or flow cytometry.

Here, we present a protocol describing how to i) assemble a self-replicating vector using the CyanoGate modular cloning toolkit, ii) introduce the vector into a cyanobacterial host by conjugation, and iii) characterize transgenic cyanobacteria strains using a plate reader or flow cytometry.

Cyanobacteria are a diverse group of prokaryotic photosynthetic organisms that can be genetically modified for the renewable production of useful ...

Visualization of DNA Repair Proteins Interaction by Immunofluorescence

Mammalian cells are constantly exposed to chemicals, radiations, and naturally occurring metabolic by-products, which create specific types of DNA insults. Genotoxic agents can damage the DNA backbone, break it, or modify the chemical nature of individual bases. Following DNA insult, DNA damage response (DDR) pathways are activated and proteins involved in the repair are recruited. A plethora of factors are involved in sensing the type of damage and activating the appropriate repair response. Failure to correctly activate and recruit DDR factors can lead to genomic instability, which underlies many human pathologies including cancer. Studies of DDR proteins can provide insights into genotoxic drug response and cellular mechanisms of drug resistance.
There are two major ways of visualizing&#160;proteins in vivo: direct observation, by tagging the protein of interest with a fluorescent protein and following it by live imaging, or indirect immunofluorescence on fixed samples. While visualization of fluorescently tagged proteins allows precise monitoring over time, direct tagging in N- or C-terminus can interfere with the protein localization or function. Observation of proteins in their unmodified, endogenous version is preferred. When DNA repair proteins are recruited to the DNA insult, their concentration increases locally and they form groups, or &#8220;foci&#8221;, that can be visualized by indirect immunofluorescence using specific antibodies.
Although detection of protein foci does not provide a definitive proof of direct interaction, co-localization of proteins in cells indicates that they regroup to the site of damage and can inform of the sequence of events required for complex formation. Careful analysis of foci spatial overlap in cells expressing wild type or mutant versions of a protein can provide precious clues on functional domains important for DNA repair function. Last, co-localization of proteins indicates possible direct interactions that can be verified by co-immunoprecipitation in cells, or direct pulldown using purified proteins.

Following DNA damage, human cells activate essential repair pathways to restore the integrity of their genome. Here, we describe the method of indirect immunofluorescence as a means to detect DNA repair proteins, analyze their spatial and temporal recruitment, and help interrogate protein-protein interaction at the sites of DNA damage.

Mammalian cells are constantly exposed to chemicals, radiations, and naturally occurring metabolic by-products, which create specific types of DNA ...

Cryo-Electron Microscopic Grid Preparation for Time-Resolved Studies using a Novel Robotic System, Spotiton

The capture of short-lived molecular states triggered by the early encounter of two or more interacting particles continues to be an experimental challenge of great interest to the field of cryo-electron microscopy (cryo-EM). A few methodological strategies have been developed that support these &#34;time-resolved&#34; studies, one of which, Spotiton-a novel robotic system-combines the dispensing of picoliter-sized sample droplets with precise temporal and spatial control. The time-resolved Spotiton workflow offers a uniquely efficient approach to interrogate early structural rearrangements from minimal sample volume. Fired from independently controlled piezoelectric dispensers, two samples land and rapidly mix on a nanowire EM grid as it plunges toward the cryogen. Potentially hundreds of grids can be prepared in rapid succession from only a few microliters of a sample. Here, a detailed step-by-step protocol of the operation of the Spotiton system is presented with a focus on troubleshooting specific problems that arise during grid preparation.

The protocol presented here describes the use of Spotiton, a novel robotic system, to deliver two samples of interest onto a self-wicking, nanowire grid that mix for a minimum of 90 ms prior to vitrification in liquid cryogen.

The capture of short-lived molecular states triggered by the early encounter of two or more interacting particles continues to be an experimental ...

Sample Preparation by 3D-Correlative Focused Ion Beam Milling for High-Resolution Cryo-Electron Tomography

Cryo-electron tomography (cryo-ET) has become the method of choice for investigating cellular ultrastructure and molecular complexes in their native, frozen-hydrated state. However, cryo-ET requires that samples are thin enough to not scatter or block the incident electron beam. For thick cellular samples, this can be achieved by cryo-focused ion beam (FIB) milling. This protocol describes how to target specific cellular sites during FIB milling using a 3D-correlative approach, which combines three-dimensional fluorescence microscopy data with information from the FIB-scanning electron microscope. Using this technique, rare cellular events and structures can be targeted with high accuracy and visualized at molecular resolution using cryo-transmission electron microscopy (cryo-TEM).

Here, we present a pipeline for 3D-correlative focused ion beam milling on guiding the preparation of cellular samples for cryo-electron tomography. The 3D position of fluorescently tagged proteins of interest is first determined by cryo-fluorescence microscopy, and then targeted for milling. The protocol is suitable for mammalian, yeast, and bacterial cells.

Cryo-electron tomography (cryo-ET) has become the method of choice for investigating cellular ultrastructure and molecular complexes in their native, ...