Supported in part by R01NS064593 and R21ES016636 (K.V.). Microscopy was performed at the VCU - Department of Neurobiology &amp; Anatomy Microscopy Facility, supported, in part, with funding from NIH-NINDS Center core grant 5P30NS047463. The spinning disc confocal microscope was purchased with an NIH-NCRR award (1S10RR027957).

A. Cell Preparation
<ol>
 <li>Normal human primary fibroblasts (GM02270) were obtained from Coriell Cell Repository, Camden, New Jersey, and immortalized with hTERT 6. Cells were grown and expanded in CellStar 6-cm dishes in media (4 ml) consisting of MEM supplemented with 20% fetal bovine serum (GIBCO), non-essential/essential amino acids, vitamins, sodium pyruvate, and penicillin/streptomycin (HyClone).</li>
 <li>Human embryonic kidney 293 (HEK293) cells were obtained from American Type Culture Collection and grown and expanded in CellStar 6-cm dishes. The cells were kept in media (4 ml) consisting of DMEM supplemented with 10% fetal bovine serum, non-essential amino acids, L-glutamine, and penicillin/streptomycin.</li>
 <li>53BP1-mCherry (N-Myc-53BP1 WT pLPC-Puro; Addgene plasmid 19836) and H2B-GFP (pCLNR-H2BG; Addgene plasmid 17735) fusion gene constructs also expressing puromycin or G418 resistance genes, respectively, were transduced (fibroblasts) or transfected (HEK293) using SuperFect (Qiagene) into cells and maintained under drug selection. Maintenance of fluorescence was periodically checked.</li>
 <li>24-48 hr before image acquisition, cells were trypsinized into a single-cell suspension and seeded at a low density onto 3.5-cm FluroDish glass bottom plates.</li>
</ol>
B. Microscope Setup and Image Acquisition
This protocol was developed using the Zeiss Cell Observer SD spinning disk confocal microscope equipped with an AxioObserver Z1 stand, a dual-channel Yokagawa CSU-X1A 5000 spinning disk unit, 2 Photometrics QuantEM 512SC emCCD cameras, an HXP 120C fiber-based illuminator, 4 lasers (a Lasos 100mW multi-line Argon [458, 488, 514 nm], 50 mW 405 nm diode, 40 mW 561 nm diode, and 30 mW 635 nm diode), AOTF, a Pecon XL multiS1 stage incubation system, Zeiss incubation modules (O2 Module S, CO2 Module S, TempModule S, Heating Unit XL S) and a Prior motorized XY stage with a NanoScanZ piezo Z insert. To minimize spherical aberrations while imaging live cells supported in an aqueous medium, a C-Apochromat 63x/1.20 Water/Corr objective lens and Zeiss Immersol W immersion fluid (with a refractive index n = 1.334) were used. For 2 channel confocal imaging, a RQFT 405/488/568/647 dichroic mirror and BP525/50 (green) and BP629/62 (red) emission filters were used. The system software used was Zeiss Axiovision (ver. 4.8.2.0) with AV4 Multi-channel/Z/T, Fast Imaging, Physiology, MosaiX, Mark &amp; Find, Dual Camera, Inside 4D, Autofocus and 3-D Deconvolution modules.
<ol>
 <li>Ensure that CO2 gas is running to the CO2 Module of the incubation system. If the microscope is supported by an anti-vibration air table, turn on the air supply (or N2 gas) for the air table.</li>
 <li>Turn on the power for the microscope stand, spinning disk unit, cameras, incubation modules, HXP illuminator, motorized stage, Argon laser, and computer.</li>
 <li>After 1 min of warm-up time, turn the ignition key for the Argon laser to "On".</li>
 <li>On the Zeiss laser control panel, turn on the switches for the laser lines to be used.</li>
 <li>Switch the toggle switch for the Lasos Argon laser controller from "standby" to "laser run" and adjust the light controller to the optimal level (i.e. just below the point where the green indicator turns red).</li>
 <li>Start up the AxioVision software. Note: the user interface for AxioVision may be customized with windows and pull-down menus that are specific for the particular microscope and components that it controls. As such, each system has a potentially unique interface. Therefore, generic instructions for software manipulation are provided in subsequent steps, rather than directions for specific software windows, tabs and/or pull-down menus.</li>
 <li>Approximately 1 hr prior to imaging, in the software, locate the controls for the incubator and turn on the heating for the upper chamber and the stage plate. Set the temperature to 37 &deg;C. Turn on the CO2 control and set the level at 5%.</li>
 <li>Select the objective lens for imaging. In this study, a 63x/1.20 N.A. C-Apochromat Water/Corr objective lens was used. Note: For this lens, an immersion medium with a refractive index similar to water (Zeiss Immersol W immersion fluid) is needed.</li>
 <li>If multiple separate positions are to be sampled over a long period of time, be certain to apply a sufficient amount of immersion medium to ensure that it is carried from one position to another.</li>
 <li>Place the dish on the stage and bring the objective lens up into contact with the bottom.</li>
 <li>Using either the microscope controls or the software, direct the emitted light to the eyepieces and select the appropriate widefield filter set for the fluorescent signal of interest (for this study, "Red" filter set for 53BP1 and a "GFP" filter set for H2B).</li>
 <li>View through the ocular lenses, focus the image, and locate a suitable field of cells.</li>
 <li>Using either the software or the microscope controls, direct the emitted light away from the eyepieces to the port with the confocal spinning disc unit.</li>
 <li>In the software, turn on the appropriate laser (for this study, the 561 nm laser for 53BP1 and the 488 nm line of the Argon laser for H2B).</li>
 <li>For each channel, adjust the intensity of the laser by adjusting the acousto-optic tunable filter (AOTF) control to an appropriate level.</li>
 <li>Select the appropriate dichroic mirror (RQFT 405/488/568/647) and emission filters (BP 629/62 for 53BP1 and BP 525/50 for H2B). Open the shutter to the spinning disc unit.</li>
 <li>Select the "Live" window to display the current field of view.</li>
 <li>In the software, open the "Camera" control, select the camera to be used (if a dual camera system) and set exposure time to approximately 100 ms. Adjust the % and EM gain as necessary. Note: The 53BP1-mCherry construct appears somewhat dim. We found it helpful to increase the EM gain.</li>
 <li>In the software, open the control for the confocal spinning disc unit and adjust the spinning disc speed by entering the camera exposure time that was set to capture a suitable image (e.g. ~100 ms). Click "Set" to lock in the change.</li>
 <li>Open "multi-dimensional acquisition". Select the channel tab and load/select appropriate channels. For this study, we're using channels defined for dsRed (561 nm laser excitation and BP 629/62 filter for emission) and GFP (488 nm laser line for excitation and BP525/50 filter for emission).</li>
 <li>To ensure image registration, a common dichroic mirror (RQFT 405/488/568/647) was used for both channels. Set the software to "Autofocus". Note: Go to Tools&rarr; Settings editor to adjust the MDA settings. Ensure that adequate laser power is set ("Adequate laser power" refers to a setting which enables you to excite the appropriate fluorophore while minimizing photo-bleaching).</li>
 <li>In the MDA window, select the z-stack tab. Select "Z-stack at current focus position". Set the range for ~10 &mu;m z-stack and choose "optimal" for the number of steps to ensure Nyquist sampling through Z. Note: The exact size of the z-stack will depend on the height of your cells. Adjust accordingly.</li>
 <li>23. Click "Start" and analyze the resulting z-stack image to ensure the settings are appropriate for what you are investigating (e.g. in this study, it was important to image the entire nucleus).</li>
 <li>Select the "T" (time) tab. For our experiment with camptothecin, we set the interval between imaging timepoints to 5 min and the overall duration of the session for 1 hr for a control (non-treated cells) video. Note: to minimize photo-bleaching of the cells, the experiment should be configured such that the AOTF blanks the laser between imaging time points.</li>
 <li>For multi-point imaging of several cells in the dish, in the MDA window, select the position tab. Ensure "Apply 'setting before/after time point' per position" is checked. Select "Mark_Find". Using the "Live" view, move the dish around and select appropriate fields of view.</li>
 <li>Click "Start" in the multi-dimensional-acquisition menu to begin the experiment and record a control video (of untreated cells).</li>
 <li>After the control video, add the appropriate treatment (in this case, 10 &mu;M camptothecin). We recorded the experimental (drug-treated cells) video at 5-10 min intervals for 2-4 hr. Note: An important issue to consider is the speed with which a given effect occurs after treatment. As seen in the video, 53BP1 foci begin to from ~5 min after addition of CPT. Though a relatively easy process, adding drug to the dish manually and re-calibrating the microscope does take time. Consideration must be given to this when designing experiments.</li>
 <li>For monitoring mitosis, we set the interval to 7.5 min and recorded for 4-5 hr (the specific settings depend upon the cell line used and the length of its cell cycle). Adjustments may need to be made to prevent photo-bleaching during longer recordings.</li>
</ol>
C. Image Processing and Analysis
This protocol was developed using Volocity software (PerkinElmer). The software used to acquire this data (AxioVision) also has the ability to process and analyze images. Users are encouraged to utilize the software available to them, and consult appropriate literature regarding their application.
<ol>
 <li>Open Volocity software. Create and name a new library, and import your video files.</li>
 <li>For viewing the files, we usually find it most helpful to use the "Extended Focus" setting. This superimposes the z-stack slices and, for this protocol, allows us to visualize 53BP1 foci in different areas of the nucleus.</li>
 <li>Adjust videos as necessary. Volocity is equipped with a range of tools to improve the quality of acquired images. By ensuring the settings on the microscope were appropriate, much time can be saved in editing later on. Often, it is helpful to deconvolve your images, and adjust the brightness/contrast. Your specific editing needs will vary based on the experiment.</li>
 <li>Add a relative time stamp and scale bar.</li>
 <li>To view cells in 3-D, switch to the "3-D Opacity" setting. This permits rotation of the 3-D rendered cells in space, thus providing multiple perspectives of structures of interest within the cells. Note: It is helpful to visualize cells in different planes to determine where structures of interest travel. For example, in the "Extended Focus" setting it is difficult to discern that the 53BP1 does, in fact, dissociate from DNA during mitosis. However, this is readily apparent in 3-D.</li>
 <li>Movies and still images may be exported into a variety of file types based on user preference.</li>
</ol>
D. Representative Results
An example of 53BP1 foci formation in response to CPT is shown in Figure 1. Cells exposed to CPT form foci within 5-10 min, and maintain these foci throughout the duration of recording. As shown in Figure 2, 53BP1 dissociates from chromatin at the onset of mitosis, forming a thin haze around the condensing chromosomes. As telophase occurs and mitosis comes to an end, 53BP1 once again aggregates into distinct foci. While the HEK293 cells were not exposed to CPT, they nevertheless formed abundant, spontaneous 53BP1 repair foci generated by endogenous DNA damage. This observation allowed us to conclude that 53BP1 does not form foci during early mitosis, in line with a previous report showing a similar effect after the exposure of cells to ionizing radiation and radiomimetic drugs 5.
<img alt="Figure 1" src="/files/ftp_upload/4251/4251fig1.jpg" /> 
 Figure 1. Camptothecin (10 &mu;M) causes DSBs and 53BP1 foci formation in cycling fibroblasts within 30 min of adding the drug to the medium.
<img alt="Figure 2" src="/files/ftp_upload/4251/4251fig2.jpg" /> 
 Figure 2. 53BP1 does not form foci during mitosis until telophase/G1 in HEK293 cells.

<ol>
	<li>Botuyan, M. V., Lee, J., Ward, I. M., Kim, J. E., Thompson, J. R., Chen, J., Mer, G. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Structural+basis+for+the+methylation+state-specific+recognition+of+histone+H4-K20+by+53BP1+and+Crb2+in+DNA+repair.">Structural basis for the methylation state-specific recognition of histone H4-K20 by 53BP1 and Crb2 in DNA repair.</a> Cell. 127, 1361-1373 (2006).</li><li>Dimitrova, N., Chen, Y. C., Spector, D. L., de Lange, T. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=53BP1+promotes+non-homologous+end+joining+of+telomeres+by+increasing+chromatin+mobility.">53BP1 promotes non-homologous end joining of telomeres by increasing chromatin mobility.</a> Nature. 456, 524-528 (2008).</li><li>Feuerhahn, S., Egly, J. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Tools+to+study+DNA+repair:+what's+in+the+box.">Tools to study DNA repair: what's in the box.</a> Trends Genet. 24, 467-474 (2008).</li><li>Giunta, S., Belotserkovskaya, R., Jackson, S. P. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=DNA+damage+signaling+in+response+to+double-strand+breaks+during+mitosis.">DNA damage signaling in response to double-strand breaks during mitosis.</a> J. Cell Biol. 190, 197-207 (2010).</li><li>Giunta, S., Jackson, S. 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Ther. 8, 730-738 (2009).</li><li>Huyen, Y., Zgheib, O., Ditullio, R. A., Gorgoulis, V. G., Zacharatos, P., Petty, T. J., Sheston, E. A., Mellert, H. S., Stavridi, E. S., Halazonetis, T. D. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Methylated+lysine+79+of+histone+H3+targets+53BP1+to+DNA+double-strand+breaks.">Methylated lysine 79 of histone H3 targets 53BP1 to DNA double-strand breaks.</a> Nature. 432, 406-411 (2004).</li><li>Jackson, S. P., Bartek, J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+DNA-damage+response+in+human+biology+and+disease.">The DNA-damage response in human biology and disease.</a> Nature. 461, 1071-1078 (2009).</li><li>Kanda, T., Sullivan, K. F., Wahl, G. 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No conflicts of interest declared.

Maintenance of genomic integrity is crucial for cell survival. Failure to preserve the genome results in premature aging, carcinogenesis, or death 8. There is intense interest in discerning how the DDR functions, stemming from its importance to both basic and clinical research. Many techniques have been developed over the years to aid in the study of how cells detect and repair DNA damage. Traditional methods such as immunocytochemistry and western blotting have been mainstays of the field, though recen...

<table border="1">
 <tbody>
 <tr>
 <td>Product</td>
 <td>Company</td>
 </tr>
 <tr>
 <td>CellStar culture dishes</td>
 <td>Greiner Bio-one</td>
 </tr>
 <tr>
 <td>FluroDish glass bottom dishes</td>
 <td>World Precision Instruments, Inc.</td>
 </tr>
 <tr>
 <td>MEM media</td>
 <td>GIBCO</td>
 </tr>
 <tr>
 <td>Non-essential amino acids</td>
 <td>GIBCO</td>
 </tr>
 <tr>
 <td>Amino acids</td>
 <td>GIBCO</td>
 </tr>
 <tr>
 <td>Vitamins</td>
 <td>GIBCO</td>
 </tr>
 <tr>
 <td>Sodium Pyruvate</td>
 <td>Invitrogen</td>
 </tr>
 <tr>
 <td>Penicillin/Streptomycin </td>
 <td>HyClone</td>
 </tr>
 <tr>
 <td>Fetal Bovine Serum</td>
 <td>GIBCO</td>
 </tr>
 <tr>
 <td>N-Myc-53BP1 WT pLPC-Puro; 
 plasmid 19836</td>
 <td>Addgene</td>
 </tr>
 <tr>
 <td>pCLNR-H2BG; plasmid 17735</td>
 <td>Addgene</td>
 </tr>
 <tr>
 <td>SuperFect</td>
 <td>Qiagen</td>
 </tr>
 <tr>
 <td>Zeiss Cell Observer SD Imaging system</td>
 <td>Zeiss</td>
 </tr>
 <tr>
 <td>AxioVision (release 4.8.2)</td>
 <td>Zeiss</td>
 </tr>
 <tr>
 <td>Zeiss Immersol W Oil</td>
 <td>Zeiss</td>
 </tr>
 <tr>
 <td>Volocity software (version 6.0)</td>
 <td>PerkinElmer</td>
 </tr>
 </tbody>
</table>

two- and three-dimensional live cell imaging of dna damage response proteins

Double-strand breaks (DSBs) are the most deleterious DNA lesions a cell can encounter. If left unrepaired, DSBs harbor great potential to generate mutations and chromosomal aberrations1. To prevent this trauma from catalyzing genomic instability, it is crucial for cells to detect DSBs, activate the DNA damage response (DDR), and repair the DNA. When stimulated, the DDR works to preserve genomic integrity by triggering cell cycle arrest to allow for repair to take place or force the cell to undergo apoptosis. The predominant mechanisms of DSB repair occur through nonhomologous end-joining (NHEJ) and homologous recombination repair (HRR) (reviewed in2). There are many proteins whose activities must be precisely orchestrated for the DDR to function properly. Herein, we describe a method for 2- and 3-dimensional (D) visualization of one of these proteins, 53BP1.
The p53-binding protein 1 (53BP1) localizes to areas of DSBs by binding to modified histones3,4, forming foci within 5-15 minutes5. The histone modifications and recruitment of 53BP1 and other DDR proteins to DSB sites are believed to facilitate the structural rearrangement of chromatin around areas of damage and contribute to DNA repair6. Beyond direct participation in repair, additional roles have been described for 53BP1 in the DDR, such as regulating an intra-S checkpoint, a G2/M checkpoint, and activating downstream DDR proteins7-9. Recently, it was discovered that 53BP1 does not form foci in response to DNA damage induced during mitosis, instead waiting for cells to enter G1 before localizing to the vicinity of DSBs6. DDR proteins such as 53BP1 have been found to associate with mitotic structures (such as kinetochores) during the progression through mitosis10.
In this protocol we describe the use of 2- and 3-D live cell imaging to visualize the formation of 53BP1 foci in response to the DNA damaging agent camptothecin (CPT), as well as 53BP1's behavior during mitosis. Camptothecin is a topoisomerase I inhibitor that primarily causes DSBs during DNA replication. To accomplish this, we used a previously described 53BP1-mCherry fluorescent fusion protein construct consisting of a 53BP1 protein domain able to bind DSBs11. In addition, we used a histone H2B-GFP fluorescent fusion protein construct able to monitor chromatin dynamics throughout the cell cycle but in particular during mitosis12. Live cell imaging in multiple dimensions is an excellent tool to deepen our understanding of the function of DDR proteins in eukaryotic cells.

Watch this Scientific Journal Video about Two- and Three-Dimensional Live Cell Imaging of DNA Damage Response Proteins at JoVE.com

Two- and Three-Dimensional Live Cell Imaging of DNA Damage Response Proteins

This protocol describes a method for visualizing a DNA double-strand break signaling protein activated in response to DNA damage as well as its localization during mitosis.

Double-strand breaks (DSBs) are the most deleterious DNA lesions a cell can encounter. If left unrepaired, DSBs harbor great potential to generate ...

two-three-dimensional-live-cell-imaging-dna-damage-response

Research

JoVE Journal

Biology

14.0K 조회수.  Virginia Commonwealth University. 이 프로토콜은 DNA 손상뿐만 아니라 유사 분열 동안의 현지화에 대한 응답으로 활성화 DNA 이중 가닥 브레이크 신호 단백질을 시각화하는 방법을 설명합니다.

비디오: Two- and Three-Dimensional Live Cell Imaging of DNA Damage Response Proteins

Microfabrication of Chip-sized Scaffolds for Three-dimensional Cell cultivation

Using microfabrication technologies is a prerequisite to create scaffolds of reproducible geometry and constant quality for three-dimensional cell cultivation. These technologies offer a wide spectrum of advantages not only for manufacturing but also for different applications. The size and shape of formed cell clusters can be influenced by the exact and reproducible architecture of the microfabricated scaffold and, therefore, the diffusion path length of nutrients and gases can be controlled.1 This is unquestionably a useful tool to prevent apoptosis and necrosis of cells due to an insufficient nutrient and gas supply or removal of cellular metabolites.
Our polymer chip, called CellChip, has the outer dimensions of 2 x 2 cm with a central microstructured area. This area is subdivided into an array of up to 1156 microcontainers with a typical dimension of 300 m edge length for the cubic design (cp- or cf-chip) or of 300 m diameter and depth for the round design (r-chip).2
So far, hot embossing or micro injection moulding (in combination with subsequent laborious machining of the parts) was used for the fabrication of the microstructured chips. Basically, micro injection moulding is one of the only polymer based replication techniques that, up to now, is capable for mass production of polymer microstructures.3 However, both techniques have certain unwanted limitations due to the processing of a viscous polymer melt with the generation of very thin walls or integrated through holes. In case of the CellChip, thin bottom layers are necessary to perforate the polymer and provide small pores of defined size to supply cells with culture medium e.g. by microfluidic perfusion of the containers.
In order to overcome these limitations and to reduce the manufacturing costs we have developed a new microtechnical approach on the basis of a down-scaled thermoforming process. For the manufacturing of highly porous and thin walled polymer chips, we use a combination of heavy ion irradiation, microthermoforming and track etching. In this so called "SMART" process (Substrate Modification And Replication by Thermoforming) thin polymer films are irradiated with energetic heavy projectiles of several hundred MeV introducing so-called "latent tracks" Subsequently, the film in a rubber elastic state is formed into three dimensional parts without modifying or annealing the tracks. After the forming process, selective chemical etching finally converts the tracks into cylindrical pores of adjustable diameter.

We present two processes for the microfabrication of porous polymer chips for three-dimensional cell cultivation. The first one is hot embossing combined with a solvent vapour welding process. The second one uses a recently developed microthermoforming process combined with ion track technology leading to a significant simplification of manufacture.

Using microfabrication technologies is a prerequisite to create scaffolds of reproducible geometry and constant quality for three-dimensional cell ...

Two-photon axotomy and time-lapse confocal imaging in live zebrafish embryos

Zebrafish have long been utilized to study the cellular and molecular mechanisms of development by time-lapse imaging of the living transparent embryo. Here we describe a method to mount zebrafish embryos for long-term imaging and demonstrate how to automate the capture of time-lapse images using a confocal microscope. We also describe a method to create controlled, precise damage to individual branches of peripheral sensory axons in zebrafish using the focused power of a femtosecond laser mounted on a two-photon microscope. The parameters for successful two-photon axotomy must be optimized for each microscope. We will demonstrate two-photon axotomy on both a custom built two-photon microscope and a Zeiss 510 confocal/two-photon to provide two examples.

Zebrafish trigeminal sensory neurons can be visualized in a transgenic line expressing GFP driven by a sensory neuron specific promoter 1. We have adapted this zebrafish trigeminal model to directly observe sensory axon regeneration in living zebrafish embryos. Embryos are anesthetized with tricaine and positioned within a drop of agarose as it solidifies. Immobilized embryos are sealed within an imaging chamber filled with phenylthiourea (PTU) Ringers. We have found that embryos can be continuously imaged in these chambers for 12-48 hours. A single confocal image is then captured to determine the desired site of axotomy. The region of interest is located on the two-photon microscope by imaging the sensory axons under low, non-damaging power. After zooming in on the desired site of axotomy, the power is increased and a single scan of that defined region is sufficient to sever the axon. Multiple location time-lapse imaging is then set up on a confocal microscope to directly observe axonal recovery from injury.

Here we describe a method for mounting zebrafish embryos for long-term imaging, two-photon imaging and tissue-damage techniques, and time-lapse confocal imaging.

Zebrafish have long been utilized to study the cellular and molecular mechanisms of development by time-lapse imaging of the living transparent embryo.  ...

Imaging Mismatch Repair and Cellular Responses to DNA Damage in Bacillus subtilis

Both prokaryotes and eukaryotes respond to DNA damage through a complex set of physiological changes. Alterations in gene expression, the redistribution of existing proteins, and the assembly of new protein complexes can be stimulated by a variety of DNA lesions and mismatched DNA base pairs. Fluorescence microscopy has been used as a powerful experimental tool for visualizing and quantifying these and other responses to DNA lesions and to monitor DNA replication status within the complex subcellular architecture of a living cell. Translational fusions between fluorescent reporter proteins and components of the DNA replication and repair machinery have been used to determine the cues that target DNA repair proteins to their cognate lesions in vivo and to understand how these proteins are organized within bacterial cells. In addition, transcriptional and translational fusions linked to DNA damage inducible promoters have revealed which cells within a population have activated genotoxic stress responses. In this review, we provide a detailed protocol for using fluorescence microscopy to image the assembly of DNA repair and DNA replication complexes in single bacterial cells. In particular, this work focuses on imaging mismatch repair proteins, homologous recombination, DNA replication and an SOS-inducible protein in Bacillus subtilis. All of the procedures described here are easily amenable for imaging protein complexes in a variety of bacterial species.

Imaging Mismatch Repair and Cellular Responses to DNA Damage in Bacillus subtilis

A detailed protocol is described for imaging the real time formation of DNA repair complexes in Bacillus subtilis cells.

Both prokaryotes and eukaryotes respond to DNA damage through a complex set of physiological changes. Alterations in gene expression, the redistribution ...

Fluorescent Labeling of COS-7 Expressing SNAP-tag Fusion Proteins for Live Cell Imaging

SNAP-tag and CLIP-tag protein labeling systems enable the specific, covalent attachment of molecules, including fluorescent dyes, to a protein of interest in live cells. These systems offer a broad selection of fluorescent substrates optimized for a range of imaging instrumentation. Once cloned and expressed, the tagged protein can be used with a variety of substrates for numerous downstream applications without having to clone again. There are two steps to using this system: cloning and expression of the protein of interest as a SNAP-tag fusion, and labeling of the fusion with the SNAP-tag substrate of choice. The SNAP-tag is a small protein based on human O6-alkylguanine-DNA-alkyltransferase (hAGT), a DNA repair protein. SNAP-tag labels are dyes conjugated to guanine or chloropyrimidine leaving groups via a benzyl linker. In the labeling reaction, the substituted benzyl group of the substrate is covalently attached to the SNAP-tag. CLIP-tag is a modified version of SNAP-tag, engineered to react with benzylcytosine rather than benzylguanine derivatives. When used in conjunction with SNAP-tag, CLIP-tag enables the orthogonal and complementary labeling of two proteins simultaneously in the same cells.

SNAP-tag and CLIP-tag protein labeling systems enable the specific, covalent attachment of molecules, including fluorescent dyes, to a protein of interest in live cells. Once cloned and expressed, the tagged protein can be used with a variety of substrates for numerous downstream applications without having to clone again.

SNAP-tag and CLIP-tag protein labeling systems enable the specific, covalent attachment of molecules, including fluorescent dyes, to a protein of interest ...

Two-Photon-Based Photoactivation in Live Zebrafish Embryos

Photoactivation of target compounds in a living organism has proven a valuable approach to investigate various biological processes such as embryonic development, cellular signaling and adult physiology. In this respect, the use of multi-photon microscopy enables quantitative photoactivation of a given light responsive agent in deep tissues at a single cell resolution. As zebrafish embryos are optically transparent, their development can be monitored in vivo. These traits make the zebrafish a perfect model organism for controlling the activity of a variety of chemical agents and proteins by focused light. Here we describe the use of two-photon microscopy to induce the activation of chemically caged fluorescein, which in turn allows us to follow cell's destiny in live zebrafish embryos. We use embryos expressing a live genetic landmark (GFP) to locate and precisely target any cells of interest. This procedure can be similarly used for precise light induced activation of proteins, hormones, small molecules and other caged compounds.

Multiphoton microscopy allows control of low energy photons with deep optical penetration and reduced phototoxicity. We describe the use of this technology for live cell labeling in zebrafish embryos. This protocol can be readily adapted for photo-induction of various light-responsive molecules.

Photoactivation of target compounds in a living organism has proven a valuable approach to investigate various biological processes such as embryonic ...

Live Imaging of GFP-labeled Proteins in Drosophila Oocytes

The Drosophila oocyte has been established as a versatile system for investigating fundamental questions such as cytoskeletal function, cell organization, and organelle structure and function. The availability of various GFP-tagged proteins means that many cellular processes can be monitored in living cells over the course of minutes or hours, and using this technique, processes such as RNP transport, epithelial morphogenesis, and tissue remodeling have been described in great detail in Drosophila oocytes1,2.
The ability to perform video imaging combined with a rich repertoire of mutants allows an enormous variety of genes and processes to be examined in incredible detail. One such example is the process of ooplasmic streaming, which initiates at mid-oogenesis3,4. This vigorous movement of cytoplasmic vesicles is microtubule and kinesin-dependent5 and provides a useful system for investigating cytoskeleton function at these stages.
Here I present a protocol for time lapse imaging of living oocytes using virtually any confocal microscopy setup.

Live Imaging of GFP-labeled Proteins in Drosophila Oocytes

A protocol for live imaging of GFP-tagged proteins or autofluorescent structures in individual Drosophila oocytes is described.

The  Drosophila oocyte has been established as a versatile system for investigating  fundamental questions such as cytoskeletal function, cell ...

4D Imaging of Protein Aggregation in Live Cells

One of the key tasks of any living cell is maintaining the proper folding of newly synthesized proteins in the face of ever-changing environmental conditions and an intracellular environment that is tightly packed, sticky, and hazardous to protein stability1. The ability to dynamically balance protein production, folding and degradation demands highly-specialized quality control machinery, whose absolute necessity is observed best when it malfunctions. Diseases such as ALS, Alzheimer's, Parkinson's, and certain forms of Cystic Fibrosis have a direct link to protein folding quality control components2, and therefore future therapeutic development requires a basic understanding of underlying processes. Our experimental challenge is to understand how cells integrate damage signals and mount responses that are tailored to diverse circumstances.
The primary reason why protein misfolding represents an existential threat to the cell is the propensity of incorrectly folded proteins to aggregate, thus causing a global perturbation of the crowded and delicate intracellular folding environment1. The folding health, or "proteostasis," of the cellular proteome is maintained, even under the duress of aging, stress and oxidative damage, by the coordinated action of different mechanistic units in an elaborate quality control system3,4. A specialized machinery of molecular chaperones can bind non-native polypeptides and promote their folding into the native state1, target them for degradation by the ubiquitin-proteasome system5, or direct them to protective aggregation inclusions6-9.
In eukaryotes, the cytosolic aggregation quality control load is partitioned between two compartments8-10: the juxtanuclear quality control compartment (JUNQ) and the insoluble protein deposit (IPOD) (Figure 1 - model). Proteins that are ubiquitinated by the protein folding quality control machinery are delivered to the JUNQ, where they are processed for degradation by the proteasome. Misfolded proteins that are not ubiquitinated are diverted to the IPOD, where they are actively aggregated in a protective compartment.
Up until this point, the methodological paradigm of live-cell fluorescence microscopy has largely been to label proteins and track their locations in the cell at specific time-points and usually in two dimensions. As new technologies have begun to grant experimenters unprecedented access to the submicron scale in living cells, the dynamic architecture of the cytosol has come into view as a challenging new frontier for experimental characterization. We present a method for rapidly monitoring the 3D spatial distributions of multiple fluorescently labeled proteins in the yeast cytosol over time. 3D timelapse (4D imaging) is not merely a technical challenge; rather, it also facilitates a dramatic shift in the conceptual framework used to analyze cellular structure.
We utilize a cytosolic folding sensor protein in live yeast to visualize distinct fates for misfolded proteins in cellular aggregation quality control, using rapid 4D fluorescent imaging. The temperature sensitive mutant of the Ubc9 protein10-12 (Ubc9ts) is extremely effective both as a sensor of cellular proteostasis, and a physiological model for tracking aggregation quality control. As with most ts proteins, Ubc9ts is fully folded and functional at permissive temperatures due to active cellular chaperones. Above 30 &deg;C, or when the cell faces misfolding stress, Ubc9ts misfolds and follows the fate of a native globular protein that has been misfolded due to mutation, heat denaturation, or oxidative damage. By fusing it to GFP or other fluorophores, it can be tracked in 3D as it forms Stress Foci, or is directed to JUNQ or IPOD.

Cellular viability depends on timely and efficient management of protein misfolding. Here we describe a method for visualizing the different potential fates of a misfolded protein: refolding, degradation, or sequestration in inclusions. We demonstrate the use of a folding sensor, Ubc9ts, for monitoring proteostasis and aggregation quality control in live cells using 4D microscopy.

One of the key tasks of any living cell is maintaining the proper folding of newly synthesized proteins in the face of ever-changing environmental ...

Mouse Fetal Whole Intestine Culture System for Ex Vivo Manipulation of Signaling Pathways and Three-dimensional Live Imaging of Villus Development

Most morphogenetic processes in the fetal intestine have been inferred from thin sections of fixed tissues, providing snapshots of changes over developmental stages. Three-dimensional information from thin serial sections can be challenging to interpret because of the difficulty of reconstructing serial sections perfectly and maintaining proper orientation of the tissue over serial sections. Recent findings by Grosse et al., 2011 highlight the importance of three- dimensional information in understanding morphogenesis of the developing villi of the intestine1. Three-dimensional reconstruction of singly labeled intestinal cells demonstrated that the majority of the intestinal epithelial cells contact both the apical and basal surfaces. Furthermore, three-dimensional reconstruction of the actin cytoskeleton at the apical surface of the epithelium demonstrated that the intestinal lumen is continuous and that secondary lumens are an artifact of sectioning. Those two points, along with the demonstration of interkinetic nuclear migration in the intestinal epithelium, defined the developing intestinal epithelium as a pseudostratified epithelium and not stratified as previously thought1. The ability to observe the epithelium three-dimensionally was seminal to demonstrating this point and redefining epithelial morphogenesis in the fetal intestine. With the evolution of multi-photon imaging technology and three-dimensional reconstruction software, the ability to visualize intact, developing organs is rapidly improving. Two-photon excitation allows less damaging penetration deeper into tissues with high resolution. Two-photon imaging and 3D reconstruction of the whole fetal mouse intestines in Walton et al., 2012 helped to define the pattern of villus outgrowth2. Here we describe a whole organ culture system that allows ex vivo development of villi and extensions of that culture system to allow the intestines to be three-dimensionally imaged during their development.

Mouse Fetal Whole Intestine Culture System for Ex Vivo Manipulation of Signaling Pathways and Three-dimensional Live Imaging of Villus Development

Improved imaging technology is allowing three-dimensional imaging of organs during development. Here we describe a whole organ culture system that allows live imaging of the developing villi in the fetal mouse intestine.

Most morphogenetic processes in the fetal intestine have been inferred from thin sections of fixed tissues, providing snapshots of changes over ...

FRET Imaging in Three-dimensional Hydrogels

Imaging of F&#246;rster resonance energy transfer (FRET)&#160;is a powerful tool for examining cell biology in real-time. Studies utilizing FRET commonly employ two-dimensional (2D) culture, which does not mimic the three-dimensional (3D) cellular microenvironment. A method to perform quenched emission FRET imaging using conventional widefield epifluorescence microscopy of cells within a 3D hydrogel environment is presented. Here an analysis method for ratiometric FRET probes that yields linear ratios over the probe activation range is described. Measurement of intracellular cyclic adenosine monophosphate (cAMP) levels is demonstrated in chondrocytes under forskolin stimulation using a probe for EPAC1 activation (ICUE1) and the ability to detect differences in cAMP signaling dependent on hydrogel material type, herein a photocrosslinking hydrogel (PC-gel, polyethylene glycol dimethacrylate) and a thermoresponsive hydrogel (TR-gel). Compared with 2D FRET methods, this method requires little additional work. Laboratories already utilizing FRET imaging in 2D can easily adopt this method to perform cellular studies in a 3D microenvironment. It can further be applied to high throughput drug screening in engineered 3D microtissues. Additionally, it is compatible with other forms of FRET imaging, such as anisotropy measurement and fluorescence lifetime imaging (FLIM), and with advanced microscopy platforms using confocal, pulsed, or modulated illumination.

F&#246;rster resonance energy transfer (FRET) imaging is a powerful tool for real-time cell biology studies. Here a method for FRET imaging cells in physiologic three-dimensional (3D) hydrogel microenvironments using conventional epifluorescence microscopy is presented. An analysis for ratiometric FRET probes that yields linear ratios over the activation range is described.

Imaging of F&#246;rster resonance energy transfer (FRET)&#160;is a powerful tool for examining cell biology in real-time. Studies utilizing FRET commonly ...

Adipo-Clear: A Tissue Clearing Method for Three-Dimensional Imaging of Adipose Tissue

Adipose tissue plays a central role in energy homeostasis and thermoregulation. It is composed of different types of adipocytes, as well as adipocyte precursors, immune cells, fibroblasts, blood vessels, and nerve projections. Although the molecular control of cell type specification and how these cells interact have been increasingly delineated, a more comprehensive understanding of these adipose-resident cells can be achieved by visualizing their distribution and architecture throughout the whole tissue. Existing immunohistochemistry and immunofluorescence approaches to analyze adipose histology rely on thin paraffin-embedded sections. However, thin sections capture only a small portion of tissue; as a result, the conclusions can be biased by what portion of tissue is analyzed. We have therefore developed an adipose tissue clearing technique, Adipo-Clear, to permit comprehensive three-dimensional visualization of molecular and cellular patterns in whole adipose tissues. Adipo-Clear was adapted from iDISCO/iDISCO+, with specific modifications made to completely remove the lipid stored in the tissue while preserving native tissue morphology. In combination with light-sheet fluorescence microscopy, we demonstrate here the use of the Adipo-Clear method to obtain high-resolution volumetric images&#160;of an entire adipose tissue.

Due to the high lipid content, adipose tissue has been challenging to visualize using traditional histological methods. Adipo-Clear is a tissue clearing technique that allows robust labeling and high-resolution volumetric fluorescent imaging of adipose tissue. Here, we describe the methods for sample preparation, pretreatment, staining, clearing, and mounting for imaging.

Adipose tissue plays a central role in energy homeostasis and thermoregulation. It is composed of different types of adipocytes, as well as adipocyte ...