The authors are supported by the Swedish research council, the Swedish foundation for strategic research, the Swedish cancer society, the Swedish child cancer society, &Aring;ke Wibergs foundation and Jeanssons foundation.

1. Introducing the probe to cells
<ol>
 <li>Co-transfect cells with a FRET-based probe and a plasmid conferring resistance. Choose a transfection method that is efficient in your cell-type of interest. For U2OS cells, standard calcium phosphate transfection methods give adequate results15.</li>
 <li>Select cells in the appropriate antibiotic for at least seven days. This enriches the amount of cells expressing the probe and limits the amount of cells with toxic expression levels, or expression levels that severely affect the cell cycle.</li>
 <li>(optional) Select for stable clones.</li>
 <li>Using fixed or live cells, verify that both CFP and YFP are present in all cells at approximately the same ratio.</li>
 <li>In case some cells contain only CFP or YFP, recombination has likely occurred. Recombination can occur either during bacterial growth or after transfection of the probe and may in the latter case depend on plasmid quality. Repeat plasmid preparation and linearize the plasmid before transfection if the problem persists.</li>
</ol>
2. Monitoring ratiometric FRET
<ol>
 <li>Seed cells on glass bottom dishes, or cover slips for mounting in a chamber. Make sure that dishes/cover slips are no 1.5 (170 &mu;m thick).</li>
 <li>Mount cells in growth medium without phenol red on a motorized epifluorescence microscope with temperature control set to 37 &deg;C. Ensure pH regulation of media, either by CO2 or by using CO2-independent media such as Liebowitz-15.</li>
 <li>Focus on cells using transmitted light. Do not look on cells using fluorescent light.</li>
 <li>Start by acquiring a 12 or 16 bit image using YFP excitation (YFPex) and YFP emission (YFPem) filters and a suitable dichroic mirror. Use maximal binning available that still permits the required subcellular resolution. Using a high binning to decrease exposure time or intensity is crucial to avoid phototoxicity.</li>
 <li>Measure or estimate the background intensity and the approximate noise by measuring or estimating the rough average, minimum and maximum pixel values in an area devoid of cells. Change exposure time and neutral density filters so that the average signal in the cell is approximately the background intensity plus five to ten times the difference between max and min background intensities.</li>
 <li>Verify that images are not close to being saturated. The maximum intensity in the image should be less than half of the dynamic range (e.g. max 2000 in a 12 bit image) to allow for intensity differences when cells enter mitosis.</li>
 <li>Repeat step 2.4 to 2.6 using CFP excitation and YFP emission filters and a suitable dichroic mirror.</li>
 <li>Select multiple regions with transfected cells, excluding the regions used for point 2.4 to 2.7, and ensure that maximum intensity is less than half of the dynamic range in all cells.</li>
 <li>Start a time-lapse experiment acquiring two images every 45 min using YFPex - YFPem and CFPex - YFPem filter sets.</li>
</ol>
3. Verifying and optimizing conditions
<ol>
 <li>Open the acquired images and monitor the cell cycle time from mitosis to mitosis for cells that express the transfected probe.</li>
 <li>Compare the measured cell cycle time to published data for the cell type used. Alternatively, film untransfected cells by acquiring one image of transmitted light only (e.g. DIC or phase-contrast) every hour to get reference cell cycle times.</li>
 <li>In case the time from mitosis to mitosis corresponds to reference cell cycle timings, proceed to step 4. Alternatively, re-acquire images using harder exposure, more time-points or multiple z-levels and repeat step 3.</li>
 <li>In case the time from mitosis to mitosis does not correspond to reference cell cycle timings, film transfected cells by acquiring one image of transmitted light only (e.g. DIC or phase-contrast) every hour followed by one image at the end of the experiment to identify transfected cells.</li>
 <li>If the transfected cells still show longer cell cycle times, repeat step 1 but transfect less probe or select for stable clones that contain very low amounts of probe.</li>
 <li>If the transfected cells show normal cell cycle times in the absence of fluorescent light, repeat step 2 but lower exposure by modifying binning, neutral density filters, exposure time, and time between images.</li>
</ol>
4. Analysing FRET
<ol>
 <li>The analysis can be performed by most software dedicated to microscopy. Here we describe how to perform the analysis using the freeware ImageJ (<a href="http://rsb.info.nih.gov/ij" target="_blank">http://rsb.info.nih.gov/ij</a>) utilizing the plugin Ratio Plus (<a href="http://imagej.nih.gov/ij/plugins/ratio-plus.html" target="_blank">http://imagej.nih.gov/ij/plugins/ratio-plus.html</a>).</li>
 <li>Open the images in ImageJ. In case your images are in the format of a multicolor stack (e.g. Deltavision), separate the individual channels by first converting them to a hyperstack: Image-hyperstack-stack to hyperstack and subsequently split them to single channel stacks using: Image-hyperstack-reduce dimensionality</li>
 <li>Multiply the YFPex - YFPem stack with 3 using: Process-Math-Multiply This step is optional, as it is only intended to increase the visual clarity of the images by ensuring that ratios are not in the range between 0 and 1.</li>
 <li>In the YFPex - YFPem stack, draw a region of interest (ROI) in an area that is devoid of cells, but that is close to a cell to be measured. Different cells in the same image may require different regions, as both background and signal intensity typically is strongest in the center of an image.</li>
 <li>Add the ROI to ROI manager: Analyze-tools-ROI manager.</li>
 <li>Measure the average intensity of the ROI in both the YFPex - YFPem and the CFPex - YFPem stack: Analyze-Measure.</li>
 <li>Repeat at multiple timepoints and verify that background intensities close to the cell of interest are similar throughout the film.</li>
 <li>Set measurements to include minimal intensity: Analyze-Set measurements-Min and Max grey value.</li>
 <li>Draw a region of interest covering most of a cell and measure the minimal intensity in both the YFPex - YFPem and the CFPex - YFPem stack. Take the difference between the measured minimal intensity and the background intensity from 4.6 and divide it by two. This provides a starting estimate for a clipping value.</li>
 <li>Open Ratio Plus. Select the YFPex - YFPem and the CFPex - YFPem stack. For FRET ratio, select CFPex - YFPem as stack 1, for inverted FRET ratio visualizing probes where FRET efficiency decreases upon phosphorylation, select YFPex - YFPem as stack 1.</li>
 <li>Insert the measured background intensities and clipping values.</li>
 <li>Set the scaling of the resulting ratio stack and apply a suitable look up table (LUT) to visualize ratio changes.</li>
</ol>
5. Representative Results
Plk1 activity is first visible in the nucleus in G2 phase and peaks during mitosis. Figure 3 shows an experiment using minimal phototoxicity settings as described in section 2. Please note that this is a representative initial result and that exposure conditions or time between images can be modified to increase signal to noise ratio or temporal resolution. Figure 3D shows that a majority of the cells expressing the probe proliferate with a cell cycle time of between 20 and 25 hours, indicating that the imaging conditions and expression levels of the probe do not affect cell cycle timings. Although there is considerable noise, the trend of Plk1 activity increasing in G2 and peaking in mitosis14 is clearly visible (fig 3A). Processing the raw data, here by mean filtering presented as a kymograph (fig 3B), or quantification of the average inverted ratio (fig 3C) can enhance clarity.
<img alt="Figure 1" src="/files/ftp_upload/3410/3410fig1.jpg" /> 
Figure 1. Principle of a FRET-based probe to monitor kinase and phosphatase activities. Two fluorophores, typically CFP (blue) and YFP (green), are connected by a phospho-binding domain (orange) and a phosphorylatable sequence (yellow). Phosphorylation (red) mediates binding to the phospho-binding domain, thereby inducing a conformational change in the probe. The conformational change results in a difference in the distance or orientation between the two fluorophores, which affects the FRET efficiency between CFP and YFP. FRET can be visualized by exciting CFP and monitoring YFP emission (dotted lines).
<img alt="Figure 2" src="/files/ftp_upload/3410/3410fig2.jpg" /> 
Figure 2. Schematic outline of the experimental procedure.
<img alt="Figure 3" src="/files/ftp_upload/3410/3410fig3.jpg" /> 
Figure 3. Plk1 activity is first visible in the nucleus in G2 phase and peaks during mitosis. U2OS cells expressing a FRET-based probe monitoring Plk1 activity9,14 were filmed for 60 hr using a Deltavision Spectris Imaging system equipped with a 20x NA 0.7 air objective and a mercury lamp. Imaging conditions were selected to cause minimal phototoxicity, as outlined in section 2, using 4x binning and neutral density filters that block 99% of the incoming light. A, false color representation of inverted FRET-ratio, following one cell through 4 divisions. B, kymograph of the cell shown in A after applying a mean filter. C, quantification of the inverted FRET ratio of the cell shown in A. D, cumulative mitotic entry of 50 FRET-probe expressing cells, including divisions of daughter cells.

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We have nothing to disclose.

Monitoring FRET throughout the cell cycle requires considerations that are less crucial when assessing short-term responses to external stimuli. First, cell cycle progression is easily perturbed by stress signaling, requiring that phototoxicity is kept to a minimum. Second, all reporters can potentially affect cellular processes by titrating out kinases, phosphatases or interaction domains. The probably most straightforward way to assess if the experimental conditions are adequate is to measure the cell cycle length from...

<table border="1">
<tbody>
 <tr>
 <td>Reagent </td>
 <td>Catalogue Number </td>
 <td>Company </td>
 </tr>
 <tr>
 <td>Leibovitz L-15, no phenol red</td>
 <td>21083-027 </td>
 <td>GIBCO, by Life Technologies </td>
 </tr>
 <tr>
 <td>DMEM+Glutamax-I</td>
 <td>31966</td>
 <td>GIBCO, by Life Technologies</td>
 </tr>
 <tr>
 <td>Fetal Bovine Serum (FBS) </td>
 <td>SV30160.03</td>
 <td>HyClone </td>
 </tr>
 <tr>
 <td>0.05% Trypsin EDTA </td>
 <td>SH30236.01</td>
 <td>HyClone </td>
 </tr>
 <tr>
 <td>Penicillin-Streptomycin</td>
 <td>SV30010 </td>
 <td>HyClone </td>
 </tr>
 <tr>
 <td>DPBS </td>
 <td>14287</td>
 <td>GIBCO, by Life Technologies</td>
 </tr>
 <tr>
 <td> Puromycin </td>
 <td>P8833 </td>
 <td>Sigma-Aldrich</td>
 </tr>
</tbody>
</table>

monitoring kinase and phosphatase activities through the cell cycle by ratiometric fret

F&ouml;rster resonance energy transfer (FRET)-based reporters1 allow the assessment of endogenous kinase and phosphatase activities in living cells. Such probes typically consist of variants of CFP and YFP, intervened by a phosphorylatable sequence and a phospho-binding domain. Upon phosphorylation, the probe changes conformation, which results in a change of the distance or orientation between CFP and YFP, leading to a change in FRET efficiency (Fig 1). Several probes have been published during the last decade, monitoring the activity balance of multiple kinases and phosphatases, including reporters of PKA2, PKB3, PKC4, PKD5, ERK6, JNK7, Cdk18, Aurora B9 and Plk19. Given the modular design, additional probes are likely to emerge in the near future10. 
Progression through the cell cycle is affected by stress signaling pathways 11. Notably, the cell cycle is regulated differently during unperturbed growth compared to when cells are recovering from stress12.Time-lapse imaging of cells through the cell cycle therefore requires particular caution. This becomes a problem particularly when employing ratiometric imaging, since two images with a high signal to noise ratio are required to correctly interpret the results. Ratiometric FRET imaging of cell cycle dependent changes in kinase and phosphatase activities has predominately been restricted to sub-sections of the cell cycle8,9,13,14.
Here, we discuss a method to monitor FRET-based probes using ratiometric imaging throughout the human cell cycle. The method relies on equipment that is available to many researchers in life sciences and does not require expert knowledge of microscopy or image processing.

Watch this Scientific Journal Video about Monitoring Kinase and Phosphatase Activities Through the Cell Cycle by Ratiometric FRET at JoVE.com

Monitoring Kinase and Phosphatase Activities Through the Cell Cycle by Ratiometric FRET

FRET-based reporters are increasingly used to monitor kinase and phosphatase activities in live cells. Here we describe a method on how to use FRET-based reporters to assess cell cycle-dependent changes in target phosphorylation.

F&ouml;rster resonance energy transfer (FRET)-based reporters1 allow the assessment of endogenous kinase and phosphatase activities in living cells. Such ...

monitoring-kinase-phosphatase-activities-through-cell-cycle

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14.2K Views.  Karolinska Institutet. FRET-based reporters are increasingly used to monitor kinase and phosphatase activities in live cells. Here we describe a method on how to use FRET-based reporters to assess cell cycle-dependent changes in target phosphorylation.

Video: Monitoring Kinase and Phosphatase Activities Through the Cell Cycle by Ratiometric FRET

Drawing Blood from Rats through the Saphenous Vein and by Cardiac Puncture

Drawing blood from rodents is necessary for a large number of both in vitro and in vivo studies. Sites of blood draws are numerous in rodents: retro-orbital sinus, jugular vein, maxillary vein, saphenous vein, heart. Each technique has its advantages and disadvantages, and some are not approved any more in some countries (e.g., retro-orbital draws in Holland). A discussion of different techniques for drawing blood are available 1-3.
Here, we present two techniques for drawing blood from rats, each with its specific applications.
Blood draw from the saphenous vein, provided it is done properly, induces minimal distress in animals and does not require anesthesia. This technique allows repeated draws of small amounts of blood, such as needed for pharmacokinetic studies 4,5, determining plasma chemistry, or blood counts 6.
Cardiac puncture allows the collection of large amounts of blood from a single animal (up to 10 ml of blood can be drawn from a 150 g rat). This technique is therefore very useful as a terminal procedure when drawing blood from the saphenous would not provide a large enough sample. We use cardiac puncture when we need sufficient amounts of serum from a specific strain of rats to grow T lymphocyte lines in vitro 4-9.

Blood draws are necessary in a large number of studies, for example to study the pharmacokinetics profile of a compound. Here, we demonstrate how to draw blood from rats using two techniques: blood draw from the saphenous vein or by cardiac puncture.

Drawing blood from rodents is necessary for a large number of both in vitro and in vivo studies. Sites of blood draws are numerous in rodents: ...

Direct Detection of the Acetate-forming Activity  of the Enzyme Acetate Kinase

Acetate kinase, a member of the acetate and sugar kinase-Hsp70-actin (ASKHA) enzyme superfamily1-5, is responsible for the reversible phosphorylation of acetate to acetyl phosphate utilizing ATP as a substrate. Acetate kinases are ubiquitous in the Bacteria, found in one genus of Archaea, and are also present in microbes of the Eukarya6. The most well characterized acetate kinase is that from the methane-producing archaeon Methanosarcina thermophila7-14. An acetate kinase which can only utilize PPi but not ATP in the acetyl phosphate-forming direction has been isolated from Entamoeba histolytica, the causative agent of amoebic dysentery, and has thus far only been found in this genus15,16.
In the direction of acetyl phosphate formation, acetate kinase activity is typically measured using the hydroxamate assay, first described by Lipmann17-20, a coupled assay in which conversion of ATP to ADP is coupled to oxidation of NADH to NAD+ by the enzymes pyruvate kinase and lactate dehydrogenase21,22, or an assay measuring release of inorganic phosphate after reaction of the acetyl phosphate product with hydroxylamine23. Activity in the opposite, acetate-forming direction is measured by coupling ATP formation from ADP to the reduction of NADP+ to NADPH by the enzymes hexokinase and glucose 6-phosphate dehydrogenase24.
Here we describe a method for the detection of acetate kinase activity in the direction of acetate formation that does not require coupling enzymes, but is instead based on direct determination of acetyl phosphate consumption. After the enzymatic reaction, remaining acetyl phosphate is converted to a ferric hydroxamate complex that can be measured spectrophotometrically, as for the hydroxamate assay. Thus, unlike the standard coupled assay for this direction that is dependent on the production of ATP from ADP, this direct assay can be used for acetate kinases that produce ATP or PPi.

A method for the determination of acetate kinase activity is described. This assay utilizes a direct reaction for determining enzyme activity and kinetics of acetate kinase in the acetate-forming direction with different phosphoryl acceptors. Furthermore, this method can be utilized for assaying other acetyl phosphate or acetyl-CoA utilizing enzymes.

Acetate kinase, a member of the acetate and sugar kinase-Hsp70-actin (ASKHA) enzyme superfamily1-5, is responsible for the reversible phosphorylation of ...

Analysis of Cell Cycle Position in Mammalian Cells

The regulation of cell proliferation is central to tissue morphogenesis during the development of multicellular organisms. Furthermore, loss of control of cell proliferation underlies the pathology of diseases like cancer. As such there is great need to be able to investigate cell proliferation and quantitate the proportion of cells in each phase of the cell cycle. It is also of vital importance to indistinguishably identify cells that are replicating their DNA within a larger population. Since a cell&prime;s decision to proliferate is made in the G1 phase immediately before initiating DNA synthesis and progressing through the rest of the cell cycle, detection of DNA synthesis at this stage allows for an unambiguous determination of the status of growth regulation in cell culture experiments.
DNA content in cells can be readily quantitated by flow cytometry of cells stained with propidium iodide, a fluorescent DNA intercalating dye. Similarly, active DNA synthesis can be quantitated by culturing cells in the presence of radioactive thymidine, harvesting the cells, and measuring the incorporation of radioactivity into an acid insoluble fraction. We have considerable expertise with cell cycle analysis and recommend a different approach. We Investigate cell proliferation using bromodeoxyuridine/fluorodeoxyuridine (abbreviated simply as BrdU) staining that detects the incorporation of these thymine analogs into recently synthesized DNA. Labeling and staining cells with BrdU, combined with total DNA staining by propidium iodide and analysis by flow cytometry1 offers the most accurate measure of cells in the various stages of the cell cycle. It is our preferred method because it combines the detection of active DNA synthesis, through antibody based staining of BrdU, with total DNA content from propidium iodide. This allows for the clear separation of cells in G1 from early S phase, or late S phase from G2/M. Furthermore, this approach can be utilized to investigate the effects of many different cell stimuli and pharmacologic agents on the regulation of progression through these different cell cycle phases.
In this report we describe methods for labeling and staining cultured cells, as well as their analysis by flow cytometry. We also include experimental examples of how this method can be used to measure the effects of growth inhibiting signals from cytokines such as TGF-&beta;1, and proliferative inhibitors such as the cyclin dependent kinase inhibitor, p27KIP1. We also include an alternate protocol that allows for the analysis of cell cycle position in a sub-population of cells within a larger culture5. In this case, we demonstrate how to detect a cell cycle arrest in cells transfected with the retinoblastoma gene even when greatly outnumbered by untransfected cells in the same culture. These examples illustrate the many ways that DNA staining and flow cytometry can be utilized and adapted to investigate fundamental questions of mammalian cell cycle control.

Determining the cell cycle position of a population of cells, or understanding how signals affect proliferation, can be readily measured by flow cytometry using this protocol. We report a simple experimental approach to staining cells and quantifying their position in the cell cycle.

The regulation of cell proliferation is central to tissue morphogenesis during the development of multicellular organisms. Furthermore, loss of control of ...

Long-term Lethal Toxicity Test with the Crustacean Artemia franciscana

Our research activities target the use of biological methods for the evaluation of environmental quality, with particular reference to saltwater/brackish water and sediment. The choice of biological indicators must be based on reliable scientific knowledge and, possibly, on the availability of standardized procedures. In this article, we present a standardized protocol that used the marine crustacean Artemia to evaluate the toxicity of chemicals and/or of marine environmental matrices. Scientists propose that the brine shrimp (Artemia) is a suitable candidate for the development of a standard bioassay for worldwide utilization. A number of papers have been published on the toxic effects of various chemicals and toxicants on brine shrimp (Artemia). The major advantage of this crustacean for toxicity studies is the overall availability of the dry cysts; these can be immediately used in testing and difficult cultivation is not demanded1,2. Cyst-based toxicity assays are cheap, continuously available, simple and reliable and are thus an important answer to routine needs of toxicity screening, for industrial monitoring requirements or for regulatory purposes3. The proposed method involves the mortality as an endpoint. The numbers of survivors were counted and percentage of deaths were calculated. Larvae were considered dead if they did not exhibit any internal or external movement during several seconds of observation4. This procedure was standardized testing a reference substance (Sodium Dodecyl Sulfate); some results are reported in this work. This article accompanies a video that describes the performance of procedural toxicity testing, showing all the steps related to the protocol.

Long-term Lethal Toxicity Test with the Crustacean Artemia franciscana

This study concerns the development and standardization of a valuable methodological protocol to determine long-term (14 days) lethal toxicity exerted by chemical substances, industrial wastewater or sewage and liquid environmental samples on the saltwater crustacean, Artemia franciscana.

Our research activities target the use of biological methods for the evaluation of environmental quality, with particular reference to saltwater/brackish ...

FRET Microscopy for Real-time Monitoring of Signaling Events in Live Cells Using Unimolecular Biosensors

F&ouml;rster resonance energy transfer (FRET) microscopy continues to gain increasing interest as a technique for real-time monitoring of biochemical and signaling events in live cells and tissues. Compared to classical biochemical methods, this novel technology is characterized by high temporal and spatial resolution. FRET experiments use various genetically-encoded biosensors which can be expressed and imaged over time in situ or in vivo1-2. Typical biosensors can either report protein-protein interactions by measuring FRET between a fluorophore-tagged pair of proteins or conformational changes in a single protein which harbors donor and acceptor fluorophores interconnected with a binding moiety for a molecule of interest3-4. Bimolecular biosensors for protein-protein interactions include, for example, constructs designed to monitor G-protein activation in cells5, while the unimolecular sensors measuring conformational changes are widely used to image second messengers such as calcium6, cAMP7-8, inositol phosphates9 and cGMP10-11. Here we describe how to build a customized epifluorescence FRET imaging system from single commercially available components and how to control the whole setup using the Micro-Manager freeware. This simple but powerful instrument is designed for routine or more sophisticated FRET measurements in live cells. Acquired images are processed using self-written plug-ins to visualize changes in FRET ratio in real-time during any experiments before being stored in a graphics format compatible with the build-in ImageJ freeware used for subsequent data analysis. This low-cost system is characterized by high flexibility and can be successfully used to monitor various biochemical events and signaling molecules by a plethora of available FRET biosensors in live cells and tissues. As an example, we demonstrate how to use this imaging system to perform real-time monitoring of cAMP in live 293A cells upon stimulation with a &beta;-adrenergic receptor agonist and blocker.

F&#246;rster resonance energy transfer (FRET) microscopy is a powerful technique for real-time monitoring of signaling events in live cells using various biosensors as reporters. Here we describe how to build a customized epifluorescence FRET imaging system from commercially available components and how to use it for FRET experiments.

F&ouml;rster resonance energy transfer (FRET) microscopy  continues to gain increasing interest as a technique for real-time monitoring  of biochemical ...

Studying DNA Looping by Single-Molecule FRET

Bending of double-stranded DNA (dsDNA) is associated with many important biological processes such as DNA-protein recognition and DNA packaging into nucleosomes. Thermodynamics of dsDNA bending has been studied by a method called cyclization which relies on DNA ligase to covalently join short sticky ends of a dsDNA. However, ligation efficiency can be affected by many factors that are not related to dsDNA looping such as the DNA structure surrounding the joined sticky ends, and ligase can also affect the apparent looping rate through mechanisms such as nonspecific binding. Here, we show how to measure dsDNA looping kinetics without ligase by detecting transient DNA loop formation by FRET (Fluorescence Resonance Energy Transfer). dsDNA molecules are constructed using a simple PCR-based protocol with a FRET pair and a biotin linker. The looping probability density known as the J factor is extracted from the looping rate and the annealing rate between two disconnected sticky ends. By testing two dsDNAs with different intrinsic curvatures, we show that the J factor is sensitive to the intrinsic shape of the dsDNA.

This study presents a detailed experimental procedure to measure looping dynamics of double-stranded DNA using single-molecule Fluorescence Resonance Energy Transfer (FRET). The protocol also describes how to extract the looping probability density called the J factor.

Bending of double-stranded DNA (dsDNA) is associated with many important biological processes such as DNA-protein recognition and DNA packaging into ...

Synchronization of Caulobacter Crescentus for Investigation of the Bacterial Cell Cycle

The cell cycle is important for growth, genome replication, and development in all cells. In bacteria, studies of the cell cycle have focused largely on unsynchronized cells making it difficult to order the temporal events required for cell cycle progression, genome replication, and division. Caulobacter crescentus provides an excellent model system for the bacterial cell cycle whereby cells can be rapidly synchronized in a G0 state by density centrifugation. Cell cycle synchronization experiments have been used to establish the molecular events governing chromosome replication and segregation, to map a genetic regulatory network controlling cell cycle progression, and to identify the establishment of polar signaling complexes required for asymmetric cell division. Here we provide a detailed protocol for the rapid synchronization of Caulobacter NA1000 cells. Synchronization can be performed in a large-scale format for gene expression profiling and western blot assays, as well as a small-scale format for microscopy or FACS assays. The rapid synchronizability and high cell yields of Caulobacter make this organism a powerful model system for studies of the bacterial cell cycle.

Synchronization of Caulobacter Crescentus for Investigation of the Bacterial Cell Cycle

Synchronization of bacterial cells is essential for studies of the bacterial cell cycle and development. Caulobacter crescentus is synchronizable through density centrifugation allowing a rapid and powerful tool for studies of the bacterial cell cycle. Here we provide a detailed protocol for the synchronization of Caulobacter cells.

The cell cycle is important for growth, genome replication, and development in all cells.  In bacteria, studies of the cell cycle have focused largely on ...

Experimental Approaches to Study Mitochondrial Localization and Function of a Nuclear Cell Cycle Kinase, Cdk1

Although mitochondria possess their own transcriptional machinery, merely 1% of mitochondrial proteins are synthesized inside the organelle. The nuclear-encoded proteins are transported into mitochondria guided by their mitochondria targeting sequences (MTS); however, a majority of mitochondrial localized proteins lack an identifiable MTS. Nevertheless, the fact that MTS can instruct proteins to go into the mitochondria provides a valuable tool for studying mitochondrial functions of normally nuclear and/or cytoplasmic proteins. We have recently identified the cell cycle kinase CyclinB1/Cdk1 complex in the mitochondria. To specifically study the mitochondrial functions of this complex, mitochondrial overexpression and knock-down of this complex without interfering with its nuclear or cytoplasmic functions were essential. By tagging CyclinB1/Cdk1 with MTS, we were able to achieve mitochondrial overexpression of this complex to study its mitochondrial targets as well as functions. Via tagging dominant-negative Cdk1 with MTS, inhibition of Cdk1 activity was accomplished particularly in the mitochondria. Potential mitochondrial targets of CyclinB1/Cdk1 complex were identified using a gel-based proteomics approach. Unlike traditional 2D gel analysis, we employed 2-dimensional difference gel electrophoresis (2D-DIGE) technology followed by phosphoprotein staining to fluorescently label differentially phosphorylated proteins in mitochondrial Cdk1 expressing cells. Identification of phosphoprotein spots that were altered in wild type versus dominant negative Cdk1 bearing mitochondria revealed the identity of mitochondrial targets of Cdk1. Finally, to determine the effect of CyclinB1/Cdk1 mitochondrial localization in cell cycle progression, a cell proliferation assay using a synthetic thymidine analogue EdU (5-ethynyl-2&#8242;-deoxyuridine) was used to monitor the cells as they go through the cell cycle and replicate their DNA. Altogether, we demonstrated a variety of approaches available to study mitochondrial localization and activity of a cell cycle kinase. These are advanced, yet easy to follow methods that will be beneficial to many cell biology researchers.

Here, we outline how to study mitochondrial localization of a (cell cycle) kinase, and how to determine its sub-mitochondrial location as well as potential mitochondrial substrates/targets. Forced expression of proteins into the mitochondria provides a useful tool for studying the functional consequences of mitochondrial localization of a protein of interest.

Although mitochondria possess their own transcriptional machinery, merely 1% of mitochondrial proteins are synthesized inside the organelle. The ...

Fecal Glucocorticoid Analysis: Non-invasive Adrenal Monitoring in Equids

Adrenal activity can be assessed in the equine species by analysis of feces for corticosterone metabolites. During a potentially aversive situation, corticotrophin releasing hormone (CRH) is released from the hypothalamus in the brain. This stimulates the release of adrenocorticotrophic hormone (ACTH) from the pituitary gland, which in turn stimulates release of glucocorticoids from the adrenal gland. In horses the glucocorticoid corticosterone is responsible for several adaptations needed to support equine flight behaviour and subsequent removal from the aversive situation. Corticosterone metabolites can be detected in the feces of horses and assessment offers a non-invasive option to evaluate long term patterns of adrenal activity. Fecal assessment offers advantages over other techniques that monitor adrenal activity including blood plasma and saliva analysis. The non-invasive nature of the method avoids sampling stress which can confound results. It also allows the opportunity for repeated sampling over time and is ideal for studies in free ranging horses. This protocol describes the enzyme linked immunoassay (EIA) used to assess feces for corticosterone, in addition to the associated biochemical validation.

Adrenal activity can be assessed in the equine species by analysis of feces for corticosterone metabolites. The method offers a non-invasive option to assess long term patterns in both domestic and free ranging horses. This protocol describes the enzyme linked immunoassay involved and the associated biochemical validation.

Adrenal activity can be assessed in the equine species by analysis of feces for corticosterone metabolites. During a potentially aversive situation, ...

G Protein-selective GPCR Conformations Measured Using FRET Sensors in a Live Cell Suspension Fluorometer Assay

F&#1255;rster resonance energy transfer (FRET)-based studies have become increasingly common in the investigation of GPCR signaling. Our research group developed an intra-molecular FRET sensor to detect the interaction between G&#945; subunits and GPCRs in live cells following agonist stimulation. Here, we detail the protocol for detecting changes in FRET between the &#946;2-adrenergic receptor and the G&#945;s C-terminus peptide upon treatment with 100 &#181;M isoproterenol hydrochloride as previously characterized1. Our FRET sensor is a single polypeptide consisting serially of a full-length GPCR, a FRET acceptor fluorophore (mCitrine), an ER/K SPASM (systematic protein affinity strength modulation) linker, a FRET donor fluorophore (mCerulean), and a G&#945; C-terminal peptide. This protocol will detail cell preparation, transfection conditions, equipment setup, assay execution, and data analysis. This experimental design detects small changes in FRET indicative of protein-protein interactions, and can also be used to compare the strength of interaction across ligands and GPCR-G protein pairings. To enhance the signal-to-noise in our measurements, this protocol requires heightened precision in all steps, and is presented here to enable reproducible execution.

Simple methods to detect the selective activation of G proteins by G protein-coupled receptors remain an outstanding challenge in cell signaling. Here, F&#1255;rster resonance energy transfer (FRET) biosensors have been developed by pairwise tethering a GPCR to G protein peptides to probe conformational changes at controlled concentrations in live cells.

F&#1255;rster resonance energy transfer (FRET)-based studies have become increasingly common in the investigation of GPCR signaling. Our research group ...