The authors thank Peter Rubenstein for useful discussion and technical advice and David Pellman for the original PB1996 clone. This work was supported by a grant from the March of Dimes and funding from the Ride for the Kids.

1. Cloning into the PB1996 Plasmid
<ol>
	<li>Design and order DNA primers from an oligonucleotide manufacturing company that flank the target sequence and contain unique restriction sites in the selected parent plasmid. NOTE: In this case, primers were designed to amplify the 400 base pairs at the 5&#8217; end of the Aip1 sequence. The XhoI restriction site was incorporated into the primer about 20 bp before the Aip1 sequence, and XmaI was included about 20 bp after the target Aip1 sequence. The plasmid used for cloning was PB1996, which contains a 3XGFP tag for fluorescence, a URA gene for yeast strain selection, and an ampicillin resistance gene for bacterial selection.</li>
	<li>Purify the genomic DNA from a Ura- yeast haploid strain. Note: Refer to Figure 1 for a diagram of cloning process.</li>
	<li>Run a standard PCR reaction using the two primers and the yeast genomic DNA to amplify the Aip1p gene with the restriction sites on either end. Digest the PCR product and the PB1996 plasmid with the restriction enzymes in separate reactions per manufacturer recommended protocols. In this case XmaI and XhoI were used with the accompanying buffer at 37 &#176;C for 1 hr.</li>
	<li>Run a 0.8% agarose electrophoresis gel to separate the digested DNA fragments. In one well, load a low DNA mass ladder and, in separate wells, load the whole reaction from each digestion. Run the gel at 100 volts for about 1 hr, until the dye front is near the end of the gel. Visualize the gel under a UV light and purify out the segment that corresponds to the protein segment and the cut plasmid.</li>
	<li>Ligate the digested Aip1 segment and PB1996 plasmid. Transform the product into DH5&#945; cells using manufacturer protocols (10 &#956;l DNA with 100 &#956;l cells for 30 min on ice, heat shock at 42 &#176;C for 2 min, grow in SOC media for 2 hr at 37 &#176;C) and plate onto culture plates that include the antibiotic for selection, in this case LB+Amp plates. Select and grow one colony in liquid culture. Purify the newly designed plasmid.</li>
</ol>
2. Mutant Yeast Strain Generation
<ol>
	<li>Use a haploid yeast strain in which the actin allele has been deleted to build a mutant yeast strain. Note: In this case, the parent strain was a generous gift from Dr. Peter Rubenstein (University of Iowa) and the construction was described by Cook et al.20 This strain lacks the chromosomal actin gene and contains a plasmid that encodes wild type yeast actin and uracil.</li>
	<li>Use the pRS plasmid encoding wild type yeast actin with a Trp+ selection as the base plasmid for mutagenesis21.</li>
	<li>Perform site-directed mutagenesis on the pRS plasmid to engineer the missense mutation into actin.
	<ol>
		<li>For the R256H mutation in actin, make the primer 5&#8217; ggtaacgaaagattccatgccccagaagc 3&#8217; to change the Arg256 to His256. Run a standard mutagenesis PCR reaction with the plasmid from step 2.2.</li>
		<li>Transform the mutant actin plasmid from the PCR into DH5&#945; cells. Isolate the plasmid DNA and sequence the actin gene to verify the mutation.</li>
	</ol>
	</li>
	<li>Transform the plasmid encoding mutant yeast actin into the haploid yeast strain from step 2.1.</li>
	<li>Culture the transformed cells on Trp- plates to select for yeast containing the Trp+ containing mutant yeast actin plasmid.</li>
	<li>Select for cells lacking the original plasmid that encodes wild type actin and uracil (described in 2.1) by sub-culturing cells from the Trp- plates on plates that contain 5-Fluoroorotic Acid (5-FOA). 5-FOA is converted to the toxic form, 5-flurouracil, in yeas strains expressing the functional URA3 gene. The cells that grow after the stepwise selection will contain only the mutant actin plasmid.</li>
	<li>Purify the plasmid DNA from the new yeast strain. Sequence the plasmid to ensure the mutant actin gene is present. Use this strain in further studies. This process is diagrammed in Figure 2.</li>
</ol>
3. Transforming the Plasmid into Yeast Cells
<ol>
	<li>Digest the Aip1p-GFP plasmid with a restriction enzyme to allow integration into the yeast chromosome. NOTE: The restriction site must be outside of the sequence block that contains the fluorescent tag, gene of interest, and selection marker. In this case, use 1 &#956;l of HpaI restriction enzyme with 8 &#956;l DNA and 1 &#956;l of the accompanying 10x buffer for 1 hr at 37 &#176;C.</li>
	<li>Culture S. cerevisiae cells (from step 2.5) that are unable to synthesize uracil (Ura- strain) overnight in 5 ml in YPD media (1% yeast extract, 2% peptone, and 2% dextrose) on a wheel at 30 &#176;C. Spin down 1 ml of the cells for 5 min at 1,000 x g.</li>
	<li>Make PLATE mixture. Make 10XTE by adding 5 ml of 1 M Tris pH 7.5 and 2 ml of 250 mM ethylenediaminetetraacetic acid (EDTA) to 43 ml of ddH2O. Add 0.204 g of lithium acetate to 20 ml of 1XTE, then 8 g of polyethylene glycol (PEG) (MW ~3,300) and filter sterilize.</li>
	<li>Decant the supernatant from the cells spun down in step 3.2 and resuspend the cells in the remaining liquid. To the suspended cells, add 2 &#956;l of 10 mg/ml salmon testes carrier DNA. Add 10 &#956;l of the digested plasmid DNA from step 3.1 and vortex. Add 0.5 ml of PLATE and vortex. Add 20 &#956;l of 1 M DTT and vortex.</li>
	<li>Incubate the mixture for 6-8 hr at 25 &#176;C. Heat shock the cells in a water bath at 42 &#176;C for 10 min. Plate 100 &#956;l of cells from the bottom of the microcentrifuge tube where they have settled out onto a &#8211;Ura plate. Incubate at 30 &#176;C for 2-3 days or until colonies form.</li>
	<li>Select individual colonies and streak out onto a &#8211;Ura plate. NOTE:&#160;These cells contain the plasmid with the Aip1-GFP and Ura gene integrated into the chromosome, allowing for growth. These cells will then be used in microscopy.</li>
</ol>
4. Visualizing the Aip1p Protein Movement Using Total Internal Reflection Fluorescence Microscopy
<ol>
	<li>Grow a culture of the yeast cells with the incorporated Aip1p-GFP in YPD overnight on a wheel at 30 &#176;C. Subculture 1 ml of the overnight culture into 9 ml of &#8211;Ura media. Grow the cells for an additional 3-4 hr on a shaker at 30 &#176;C to ensure that they are in log growth phase.</li>
	<li>Add 3 &#956;l of cells to a glass microscope slide and add a coverslip. Allow the cells to settle on the slide for 5 min. Place the slide in the bracket of the stage plate.</li>
	<li>Observe the Aip1p-GFP protein with a total internal reflection fluorescence (TIRF) microscope (488 nm) using an oil immersion 100X TIRF objective and a digital CCD camera. Using SlideBook 5.0 software, obtain images for 20 sec at a rate of 5 frames/sec (Individual frames of acquired images are presented in Figure 2A).
	<ol>
		<li>To focus on the cells, open SlideBook 5.0 software and click the Focus Window button in the toolbar to access the focus controls. Select the Scope tab and select the 100X-TIRF objective. Turn the lamp on and slide the lamp bar to 15%. Change the Bin setting to 2 x 2. Change filter to bright field.</li>
		<li>On the microscope, use the fine focus dial to bring the cells into focus as seen on the computer screen.</li>
		<li>Using the controls within the Focus Window, turn the lamp off, change the filter to Live, and click the TIRFM button.</li>
		<li>On the TIRFM illuminator attached to the right side port of the microscope, turn the laser emission to On.</li>
		<li>Select the Stream tab within the Focus Window, check the box next to Start Recording and then click bright field button.</li>
		<li>On the TIRFM illuminator, turn the micrometer to adjust the incident angle of the laser. This is used to optimize the fluorescence emission from the Aip1-GFP foci and minimize the background fluorescence from the cells and slide.</li>
		<li>When the desired image is displayed, press Start. After 20 sec, press Stop. Save the images. Under the File tab on the main menu bar, select Save Slide As and enter the file location and name.</li>
	</ol>
	</li>
	<li>Use ImageJ software to analyze the images. Use the macro plugin &#8220;Manual Tracker&#8221; to track the movement and rate of the fluorescent Aip1p foci. Change the time interval parameter to 0.2 sec.
	<ol>
		<li>Using the add track selection, select and track an individual fluorescent protein through four to eight connective frames. Use the end track command and the velocity of protein movement between each frame will display in a results window.</li>
		<li>Determine the mean rate between each frame using a Microsoft Excel spreadsheet. Use the values to compute the average velocity of Aip1p movement for each cell strain of interest. NOTE: A scatter plot of the rate of Aip1p movement and the average non-linear velocity is shown in Figure 2B.</li>
	</ol>
	</li>
</ol>

<ol>
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J., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Segregation+of+a+missense+variant+in+enteric+smooth+muscle+actin+gamma-2+with+autosomal+dominant+familial+visceral+myopathy.">Segregation of a missense variant in enteric smooth muscle actin gamma-2 with autosomal dominant familial visceral myopathy.</a> Gastroenterology. 143, 1482-1491 (2012).</li><li>Matsson, H., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Alpha-cardiac+actin+mutations+produce+atrial+septal+defects.">Alpha-cardiac actin mutations produce atrial septal defects.</a> Hum Mol Genet. 17, 256-265 (2008).</li><li>Zhu, M., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Mutations+in+the+gamma-actin+gene+(ACTG1)+are+associated+with+dominant+progressive+deafness+(DFNA20/26).">Mutations in the gamma-actin gene (ACTG1) are associated with dominant progressive deafness (DFNA20/26).</a> Am J Hum Genet. 73, 1082-1091 (2003).</li><li>Nowak, K. 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The authors have nothing to disclose.

An effective strategy to visualize the dynamics of the cytoskeleton and the utility in investigations on pathogenic mutations has been described here. Advanced imaging modalities have created new opportunities to understand the intracellular movement of proteins near the cell membrane. Total internal reflection fluorescence microscopy (TIRF) is a sensitive technique for functional studies in living cells. TIRF uses an angled excitation laser that creates an evanescent field due to the difference in refractive indexes of ...

Actin is the dominant protein comprising the cytoskeleton and participates in critical cellular processes including cell division, organelle movement, cell motility, contraction, and signaling. Over the past decade, disease-causing mutations in actin have been discovered in each of the six human actin isoforms leading to a range of disorders, from myopathies to coronary artery disease1-7. The processes by which actin mutations lead to disease continue to be elucidated. The yeast model remains the gold standard to study the biochemical effects of mutations on actin function owing to the advantages of the single essential actin isoform, genetic tractability and high conservation of actin sequence and function. Studies show that individual actin mutations lead to molecular specific dysfunctions with dominant negative effects8. For example, deafness-causing mutations in &#947;-non-muscle actin that affect the Lys-118 residue alter regulation by the actin binding protein Arp2/39. Studies frequently employ in vitro analyses of protein: protein interactions. Investigations into the effect of actin mutations on cell biology and, in particular, actin binding protein localization in the cell are limited.
Studies of the yeast cytoskeleton in vivo conventionally rely on images of fixed cells from an inverted fluorescence microscope10. These experiments supplied foundational data about the morphology of the actin cytoskeleton. Investigations have since incorporated three dimensional confocal imaging to visualize the complex cytoskeletal network11,12. This imaging permits quantification of the abundance and relative location of actin patches and filaments. Thin section electron tomography has been used to image the morphology of the dense filamentous networks relative to preserved subcellular structures13. Crowded cellular spaces with a small cross section can be examined in fine detail with this technique. Imaging studies have been extended to living cells using time lapse fluorescent microscopy. When photo bleaching and background fluorescence can be moderated, time lapse imaging allows investigations as to the dynamics of cytoskeletal proteins and the response to environmental conditions11,14. Separately, visualization of the dynamics of actin filaments in vitro was advanced by the introduction of total internal reflection fluorescence (TIRF) microscopy. Compared to wide field microscopy, TIRF has the advantage of decreased background fluorescence and enhanced contrast to monitor individual filaments15,16. With these qualities, TIRF microscopy has been adapted by cell biologists to monitor cellular structures at the plasma membrane17,18. Cellular events, including changes in the cytoskeleton, can be visualized real time with low phototoxicity, maximal contrast, and minimal background florescence19.
To better understand the effect of actin mutations on the movement, localization, and turnover of cytoskeletal proteins in the cell, TIRF microscopy and protein tagging were used. Herein, methods to study the effects of a clinically relevant mutation in actin on cytoskeletal dynamics in Saccharomyces cerevisiae are described. Specifically, the localization and movement of the actin binding protein, Aip1p, was visualized and quantified in cells expressing the R256H mutation in actin. These techniques complement in vitro biochemical studies and allow for a greater understanding of protein interactions and functions.

<table border="0" cellpadding="0" cellspacing="0" width="600">
	<colgroup>
	</colgroup>
	<tbody>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">Agarose</td>
			<td style="width:103px;">rpi</td>
			<td style="width:119px;">9012-36-6</td>
			<td style="width:167px;"></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">Bromophenol Blue</td>
			<td style="width:103px;">Amresco</td>
			<td style="width:119px;">115-39-9</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">BSA</td>
			<td style="width:103px;">NEB</td>
			<td style="width:119px;">B9001S</td>
			<td></td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;width:216px;">Change-IT Multiple Mutation Site Directed Mutagenesis Kit</td>
			<td style="width:103px;">USB Corporation</td>
			<td align="right" style="width:119px;">4166059</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">CutSmart Buffer</td>
			<td style="width:103px;">NEB</td>
			<td style="width:119px;">B7204S</td>
			<td></td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;width:216px;">DNA, single stranded from salmon testes</td>
			<td style="width:103px;">Sigma</td>
			<td style="width:119px;">9007-49-2</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">EDTA pH 7.4</td>
			<td style="width:103px;">Sigma</td>
			<td align="right" style="width:119px;">93302</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">Ethidium Bromide</td>
			<td style="width:103px;">Invitrogen</td>
			<td style="width:119px;">15585-011</td>
			<td>Warning! Harmful irritation</td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;width:216px;">Fungal/Bacterial DNA Kit</td>
			<td style="width:103px;">Symo Research</td>
			<td style="width:119px;">D6005</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">HpaI</td>
			<td style="width:103px;">NEB</td>
			<td style="width:119px;">R0105S</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">Lithium Acetate</td>
			<td style="width:103px;">AlfaAesar</td>
			<td style="width:119px;">6108-17-4</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">Low DNA Mass Ladder</td>
			<td style="width:103px;">Invitrogen</td>
			<td style="width:119px;">10068-013</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">NE Buffer #4</td>
			<td style="width:103px;">NEB</td>
			<td style="width:119px;">B7004S</td>
			<td></td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;width:216px;">Platinum PCR SuperMix High Fidelity</td>
			<td style="width:103px;">Invitrogen</td>
			<td style="width:119px;">12532-016</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">Miniprep Kit</td>
			<td style="width:103px;">Qiagen</td>
			<td align="right" style="width:119px;">27106</td>
			<td>Any kit will work</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">Quick Ligation Kit</td>
			<td style="width:103px;">NEB</td>
			<td style="width:119px;">M2200S</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">Sodium azide</td>
			<td style="width:103px;">Sigma</td>
			<td style="width:119px;">26628-22-8</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">PBS</td>
			<td style="width:103px;">Invitrogen</td>
			<td style="width:119px;">10010-023</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">PEG</td>
			<td style="width:103px;">Amresco</td>
			<td style="width:119px;">25322-68-3</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">Tris Base Ultrapure</td>
			<td style="width:103px;">rpi</td>
			<td style="width:119px;">77-86-1</td>
			<td></td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;width:216px;">Wizard SV Gel and PCR Clean-Up System</td>
			<td style="width:103px;">Promega</td>
			<td align="right" style="width:119px;">1/6/2015</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">XhoI</td>
			<td style="width:103px;">NEB</td>
			<td style="width:119px;">R0146S</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">XmaI</td>
			<td style="width:103px;">NEB</td>
			<td style="width:119px;">R0180S</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">YPD media</td>
			<td style="width:103px;">LabExpress</td>
			<td align="right" style="width:119px;">3011</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">-URA Media</td>
			<td style="width:103px;">LabExpress</td>
			<td align="right" style="width:119px;">3010</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">PCR Machine</td>
			<td style="width:103px;">Invitrogen</td>
			<td align="right" style="width:119px;">4359659</td>
			<td>Any PCR machine will work</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">TIRF Microscope</td>
			<td style="width:103px;">Olympus IX81</td>
			<td style="width:119px;"></td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">Hamamatsu ORCA-R camera</td>
			<td style="width:103px;">Hamamatsu</td>
			<td style="width:119px;"></td>
			<td></td>
		</tr>
	</tbody>
</table>

aip1p dynamics are altered by the r256h mutation in actin

Mutations in actin cause a range of human diseases due to specific molecular changes that often alter cytoskeletal function. In this study, imaging of fluorescently tagged proteins using total internal fluorescence (TIRF) microscopy is used to visualize and quantify changes in cytoskeletal dynamics. TIRF microscopy and the use of fluorescent tags also allows for quantification of the changes in cytoskeletal dynamics caused by mutations in actin. Using this technique, quantification of cytoskeletal function in live cells valuably complements in vitro studies of protein function. As an example, missense mutations affecting the actin residue R256 have been identified in three human actin isoforms suggesting this amino acid plays an important role in regulatory interactions. The effects of the actin mutation R256H on cytoskeletal movements were studied using the yeast model. The protein, Aip1, which is known to assist cofilin in actin depolymerization, was tagged with green fluorescent protein (GFP) at the N-terminus and tracked in vivo using TIRF microscopy. The rate of Aip1p movement in both wild type and mutant strains was quantified. In cells expressing R256H mutant actin, Aip1p motion is restricted and the rate of movement is nearly half the speed measured in wild type cells (0.88 &#177; 0.30 &#956;m/sec in R256H cells compared to 1.60 &#177; 0.42 &#956;m/sec in wild type cells, p &#60; 0.005).

A method to image the dynamics of cytoskeletal proteins in the cell is presented. The actin-binding protein, Aip1p, was tagged with GFP. The design for the plasmid encoding the tagged product is shown in Figure 1. The plasmid was then transformed into the yeast cells. Expression of the fluorescently tagged Aip1p allowed visualization of the protein behavior in the cell. Aip1p typically localizes to actin patches at sites of endocytosis22. To quantify Aip1p movement, more than 50 fluorescent Ai...

Watch this Scientific Journal Video about Aip1p Dynamics Are Altered by the R256H Mutation in Actin at JoVE.com

Aip1p Dynamics Are Altered by the R256H Mutation in Actin

Disease-causing mutations in actin can alter cytoskeletal function. Cytoskeletal dynamics are quantified through imaging of fluorescently tagged proteins using total internal fluorescence microscopy. As an example, the cytoskeletal protein, Aip1p, has altered localization and movement in cells expressing the mutant actin isoform, R256H.

Mutations in actin cause a range of human diseases due to specific molecular changes that often alter cytoskeletal function. In this study, imaging of ...

aip1p-dynamics-are-altered-by-the-r256h-mutation-in-actin

Research

JoVE Journal

Biology

7.9K Views.  University of Iowa.  Malattia mutazioni che causano in actina possono alterare la funzione del citoscheletro. Dinamica del citoscheletro vengono quantificati attraverso l'imaging di proteine ​​con targhetta modo fluorescente utilizzando totale microscopia a fluorescenza interna. Come esempio, la proteina del citoscheletro, Aip1p, ha modificato la localizzazione e il movimento in cellule esprimenti l'isoforma actina mutante, R256H. 

Video: Aip1p Dynamics Are Altered by the R256H Mutation in Actin

Actin Co-Sedimentation Assay; for the Analysis of Protein Binding to F-Actin

The actin cytoskeleton within the cell is a network of actin filaments that allows the movement of cells and cellular processes, and that generates tension and helps maintains cellular shape.  Although the actin cytoskeleton is a rigid structure, it is a dynamic structure that is constantly remodeling.  A number of proteins can bind to the actin cytoskeleton.  The binding of a particular protein to F-actin is often desired to support cell biological observations or to further understand dynamic processes due to remodeling of the actin cytoskeleton. The actin co-sedimentation assay is an in vitro assay routinely used to analyze the binding of specific proteins or protein domains with F-actin.  The basic principles of the assay involve an incubation of the protein of interest (full length or domain of) with F-actin, ultracentrifugation step to pellet F-actin and analysis of the protein co-sedimenting with F-actin.  Actin co-sedimentation assays can be designed accordingly to measure actin binding affinities and in competition assays.

Proteins bind to filamentous actin (F-actin) through distinct actin binding modules. In this video we demonstrate the procedure of actin co-sedimentation, which is an in vitro assay routinely used to analyze proteins or specific domains that bind F-actin.

The actin cytoskeleton within the cell is a network of actin filaments that allows the movement of cells and cellular processes, and that generates ...

Live Cell Imaging of F-actin Dynamics via Fluorescent Speckle Microscopy (FSM)

In this protocol we describe the use of Fluorescent Speckle Microscopy (FSM) to capture high-resolution images of actin dynamics in PtK1 cells. A unique advantage of FSM is its ability to capture the movement and turnover kinetics (assembly/disassembly) of the F-actin network within living cells. This technique is particularly useful in deriving quantitative measurements of F-actin dynamics when paired with computer vision software (qFSM).  We describe  the selection, microinjection and visualization of fluorescent actin probes in living cells. Importantly, similar procedures are applicable to visualizing other macomolecular assemblies. FSM has been demonstrated for microtubules, intermediate filaments, and adhesion complexes.  

Live Cell Imaging of F-actin Dynamics via Fluorescent Speckle Microscopy FSM

Selection, microinjection, and imaging of fluorescently-labeled F-actin via fluorescent speckle microscopy (FSM).

In this protocol we describe the use of Fluorescent Speckle Microscopy (FSM) to capture high-resolution images of actin dynamics in PtK1 cells. A unique ...

Tracking Dynamics of Muscle Engraftment in Small Animals by In Vivo Fluorescent Imaging

Muscular dystrophies are a group of degenerative muscle diseases characterized by progressive loss of contractile muscle cells. Currently, there is no curative treatment available. Recent advances in stem cell biology have generated new hopes for the development of effective cell based therapies to treat these diseases. Transplantation of various types of stem cells labeled with fluorescent proteins into muscles of dystrophic animal models has been used broadly in the field. A non-invasive technique with the capability to track the transplanted cell fate longitudinally can further our ability to evaluate muscle engraftment by transplanted cells more accurately and efficiently.
In the present study, we demonstrate that in vivo fluorescence imaging is a sensitive and reliable method for tracking transplanted GFP (Green Fluorescent Protein)-labeled cells in mouse skeletal muscles. Despite the concern about background due to the use of an external light necessary for excitation of fluorescent protein, we found that by using either nude mouse or eliminating hair with hair removal reagents much of this problem is eliminated. Using a CCD camera, the fluorescent signal can be detected in the tibialis anterior (TA) muscle after injection of 5 x 105 cells from either GFP transgenic mice or eGFP transduced myoblast culture. For more superficial muscles such as the extensor digitorum longus (EDL), injection of fewer cells produces a detectable signal. Signal intensity can be measured and quantified as the number of emitted photons per second in a region of interest (ROI). Since the acquired images show clear boundaries demarcating the engrafted area, the size of the ROI can be measured as well. If the legs are positioned consistently every time, the changes in total number of photons per second per muscle and the size of the ROI reflect the changes in the number of engrafted cells and the size of the engrafted area. Therefore the changes in the same muscle over time are quantifiable. In vivo fluorescent imaging technique has been used primarily to track the growth of tumorogenic cells, our study shows that it is a powerful tool that enables us to track the fate of transplanted stem cells.

Tracking Dynamics of Muscle Engraftment in Small Animals by In Vivo Fluorescent Imaging

We describe an in vivo fluorescence imaging protocol to monitor muscle regeneration by GFP-labeled myoblasts after transplantation into skeletal muscles of both healthy and dystrophic mice. This protocol can be adapted to study muscle regeneration by transplantation of other types of cells and in other muscular conditions as well.

Muscular dystrophies are a group of degenerative muscle diseases characterized by progressive loss of contractile muscle cells. Currently, there is no ...

Photobleaching Assays (FRAP &amp; FLIP) to Measure Chromatin Protein Dynamics in Living Embryonic Stem Cells

Fluorescence Recovery After Photobleaching (FRAP) and Fluorescence Loss In Photobleaching (FLIP) enable the study of protein dynamics in living cells with good spatial and temporal resolution. Here we describe how to perform FRAP and FLIP assays of chromatin proteins, including H1 and HP1, in mouse embryonic stem (ES) cells. In a FRAP experiment, cells are transfected, either transiently or stably, with a protein of interest fused with the green fluorescent protein (GFP) or derivatives thereof (YFP, CFP, Cherry, etc.). In the transfected, fluorescing cells, an intense focused laser beam bleaches a relatively small region of interest (ROI). The laser wavelength is selected according to the fluorescent protein used for fusion. The laser light irreversibly bleaches the fluorescent signal of molecules in the ROI and, immediately following bleaching, the recovery of the fluorescent signal in the bleached area - mediated by the replacement of the bleached molecules with the unbleached molecules - is monitored using time lapse imaging. The generated fluorescence recovery curves provide information on the protein's mobility. If the fluorescent molecules are immobile, no fluorescence recovery will be observed. In a complementary approach, Fluorescence Loss in Photobleaching (FLIP), the laser beam bleaches the same spot repeatedly and the signal intensity is measured elsewhere in the fluorescing cell. FLIP experiments therefore measure signal decay rather than fluorescence recovery and are useful to determine protein mobility as well as protein shuttling between cellular compartments. Transient binding is a common property of chromatin-associated proteins. Although the major fraction of each chromatin protein is bound to chromatin at any given moment at steady state, the binding is transient and most chromatin proteins have a high turnover on chromatin, with a residence time in the order of seconds. These properties are crucial for generating high plasticity in genome expression1. Photobleaching experiments are therefore particularly useful to determine chromatin plasticity using GFP-fusion versions of chromatin structural proteins, especially in ES cells, where the dynamic exchange of chromatin proteins (including heterochromatin protein 1 (HP1), linker histone H1 and core histones) is higher than in differentiated cells2,3.

Photobleaching Assays FRAP &amp; FLIP to Measure Chromatin Protein Dynamics in Living Embryonic Stem Cells

We describe photobleaching methods including Fluorescence Recovery After Photobleaching (FRAP) and Fluorescence Loss In Photobleaching (FLIP) to monitor chromatin protein dynamics in embryonic stem (ES) cells. Chromatin protein dynamics, which is considered to be one of the means to study chromatin plasticity, is enhanced in pluripotent cells.

Fluorescence Recovery After Photobleaching (FRAP) and Fluorescence Loss In Photobleaching (FLIP) enable the study of protein dynamics in living cells with ...

Study of the Actin Cytoskeleton in Live Endothelial Cells Expressing GFP-Actin

The microvascular endothelium plays an important role as a selectively permeable barrier to fluids and solutes. The adhesive junctions between endothelial cells regulate permeability of the endothelium, and many studies have indicated the important contribution of the actin cytoskeleton to determining junctional integrity1-5. A cortical actin belt is thought to be important for the maintenance of stable junctions1, 2, 4, 5. In contrast, actin stress fibers are thought to generate centripetal tension within endothelial cells that weakens junctions2-5. Much of this theory has been based on studies in which endothelial cells are treated with inflammatory mediators known to increase endothelial permeability, and then fixing the cells and labeling F-actin for microscopic observation. However, these studies provide a very limited understanding of the role of the actin cytoskeleton because images of fixed cells provide only snapshots in time with no information about the dynamics of actin structures5. 
Live-cell imaging allows incorporation of the dynamic nature of the actin cytoskeleton into the studies of the mechanisms determining endothelial barrier integrity. A major advantage of this method is that the impact of various inflammatory stimuli on actin structures in endothelial cells can be assessed in the same set of living cells before and after treatment, removing potential bias that may occur when observing fixed specimens. Human umbilical vein endothelial cells (HUVEC) are transfected with a GFP-&beta;-actin plasmid and grown to confluence on glass coverslips. Time-lapse images of GFP-actin in confluent HUVEC are captured before and after the addition of inflammatory mediators that elicit time-dependent changes in endothelial barrier integrity. These studies enable visual observation of the fluid sequence of changes in the actin cytoskeleton that contribute to endothelial barrier disruption and restoration. 
Our results consistently show local, actin-rich lamellipodia formation and turnover in endothelial cells. The formation and movement of actin stress fibers can also be observed. An analysis of the frequency of formation and turnover of the local lamellipodia, before and after treatment with inflammatory stimuli can be documented by kymograph analyses. These studies provide important information on the dynamic nature of the actin cytoskeleton in endothelial cells that can used to discover previously unidentified molecular mechanisms important for the maintenance of endothelial barrier integrity.

Microscopic imaging of live endothelial cells expressing GFP-actin allows characterization of dynamic changes in cytoskeletal structures. Unlike techniques that use fixed specimens, this method provides a detailed assessment of temporal changes in the actin cytoskeleton in the same cells before, during, and after various physical, pharmacological, or inflammatory stimuli.

The microvascular endothelium plays an important role as a selectively permeable barrier to fluids and solutes. The adhesive junctions between endothelial ...

Selective Capture of 5-hydroxymethylcytosine from Genomic DNA

5-methylcytosine (5-mC) constitutes ~2-8% of the total cytosines in human genomic DNA and impacts a broad range of biological functions, including gene expression, maintenance of genome integrity, parental imprinting, X-chromosome inactivation, regulation of development, aging, and cancer1. Recently, the presence of an oxidized 5-mC, 5-hydroxymethylcytosine (5-hmC), was discovered in mammalian cells, in particular in embryonic stem (ES) cells and neuronal cells2-4. 5-hmC is generated by oxidation of 5-mC catalyzed by TET family iron (II)/&alpha;-ketoglutarate-dependent dioxygenases2, 3. 5-hmC is proposed to be involved in the maintenance of embryonic stem (mES) cell, normal hematopoiesis and malignancies, and zygote development2, 5-10. To better understand the function of 5-hmC, a reliable and straightforward sequencing system is essential. Traditional bisulfite sequencing cannot distinguish 5-hmC from 5-mC11. To unravel the biology of 5-hmC, we have developed a highly efficient and selective chemical approach to label and capture 5-hmC, taking advantage of a bacteriophage enzyme that adds a glucose moiety to 5-hmC specifically12.
Here we describe a straightforward two-step procedure for selective chemical labeling of 5-hmC. In the first labeling step, 5-hmC in genomic DNA is labeled with a 6-azide-glucose catalyzed by &beta;-GT, a glucosyltransferase from T4 bacteriophage, in a way that transfers the 6-azide-glucose to 5-hmC from the modified cofactor, UDP-6-N3-Glc (6-N3UDPG). In the second step, biotinylation, a disulfide biotin linker is attached to the azide group by click chemistry. Both steps are highly specific and efficient, leading to complete labeling regardless of the abundance of 5-hmC in genomic regions and giving extremely low background. Following biotinylation of 5-hmC, the 5-hmC-containing DNA fragments are then selectively captured using streptavidin beads in a density-independent manner. The resulting 5-hmC-enriched DNA fragments could be used for downstream analyses, including next-generation sequencing.
Our selective labeling and capture protocol confers high sensitivity, applicable to any source of genomic DNA with variable/diverse 5-hmC abundances. Although the main purpose of this protocol is its downstream application (i.e., next-generation sequencing to map out the 5-hmC distribution in genome), it is compatible with single-molecule, real-time SMRT (DNA) sequencing, which is capable of delivering single-base resolution sequencing of 5-hmC.

Described is a two-step labeling process using &beta;-glucosyltransferase (&beta;-GT) to transfer an azide-glucose to 5-hmC, followed by click chemistry to transfer a biotin linker for easy and density-independent enrichment. This efficient and specific labeling method enables enrichment of 5-hmC with extremely low background and high-throughput epigenomic mapping via next-generation sequencing.

5-methylcytosine (5-mC) constitutes ~2-8% of the total cytosines in human genomic DNA and impacts a broad range of biological functions, including gene ...

Robust 3D DNA FISH Using Directly Labeled Probes

3D DNA FISH has become a major tool for analyzing three-dimensional organization of the nucleus, and several variations of the technique have been published. In this article we describe a protocol which has been optimized for robustness, reproducibility, and ease of use. Brightly fluorescent directly labeled probes are generated by nick-translation with amino-allyldUTP followed by chemical coupling of the dye. 3D DNA FISH is performed using a freeze-thaw step for cell permeabilization and a heating step for simultaneous denaturation of probe and nuclear DNA. The protocol is applicable to a range of cell types and a variety of probes (BACs, plasmids, fosmids, or Whole Chromosome Paints) and allows for high-throughput automated imaging. With this method we routinely investigate nuclear localization of up to three chromosomal regions.

We describe a robust and versatile protocol for analyzing nuclear architecture by 3D DNA FISH using directly labeled fluorescent probes.

3D DNA FISH has become a major tool for  analyzing three-dimensional organization of the nucleus, and several variations  of the technique have been ...

In Vivo 4-Dimensional Tracking of Hematopoietic Stem and Progenitor Cells in Adult Mouse Calvarial Bone Marrow

Through a delicate balance between quiescence and proliferation, self renewal and production of differentiated progeny, hematopoietic stem cells (HSCs) maintain the turnover of all mature blood cell lineages. The coordination of the complex signals leading to specific HSC fates relies upon the interaction between HSCs and the intricate bone marrow microenvironment, which is still poorly understood[1-2].
We describe how by combining a newly developed specimen holder for stable animal positioning with multi-step confocal and two-photon in vivo imaging techniques, it is possible to obtain high-resolution 3D stacks containing HSPCs and their surrounding niches and to monitor them over time through multi-point time-lapse imaging. High definition imaging allows detecting ex vivo labeled hematopoietic stem and progenitor cells (HSPCs) residing within the bone marrow. Moreover, multi-point time-lapse 3D imaging, obtained with faster acquisition settings, provides accurate information about HSPC movement and the reciprocal interactions between HSPCs and stroma cells.
Tracking of HSPCs in relation to GFP positive osteoblastic cells is shown as an exemplary application of this method. This technique can be utilized to track any appropriately labeled hematopoietic or stromal cell of interest within the mouse calvarium bone marrow space.

In Vivo 4-Dimensional Tracking of Hematopoietic Stem and Progenitor Cells in Adult Mouse Calvarial Bone Marrow

The nature of the interactions between hematopoietic stem and progenitor cells (HSPCs) and bone marrow niches is poorly understood. Custom hardware modifications and a multi-step acquisition protocol allow the use of two-photon and confocal microscopy to image ex vivo labeled HSPCs homed within bone marrow areas, tracking interactions and movement.

Through a delicate balance between quiescence and proliferation, self renewal and production of differentiated progeny, hematopoietic stem cells (HSCs) ...

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, ...

High-resolution Time-lapse Imaging and Automated Analysis of Microtubule Dynamics in Living Human Umbilical Vein Endothelial Cells

The physiological process by which new vasculature forms from existing vasculature requires specific signaling events that trigger morphological changes within individual endothelial cells (ECs). These processes are critical for homeostatic maintenance such as wound healing, and are also crucial in promoting tumor growth and metastasis. EC morphology is defined by the organization of the cytoskeleton, a tightly regulated system of actin and microtubule (MT) dynamics that is known to control EC branching, polarity and directional migration, essential components of angiogenesis. To study MT dynamics, we used high-resolution fluorescence microscopy coupled with computational image analysis of fluorescently-labeled MT plus-ends to investigate MT growth dynamics and the regulation of EC branching morphology and directional migration. Time-lapse imaging of living Human Umbilical Vein Endothelial Cells (HUVECs) was performed following transfection with fluorescently-labeled MT End Binding protein 3 (EB3) and Mitotic Centromere Associated Kinesin (MCAK)-specific cDNA constructs to evaluate effects on MT dynamics. PlusTipTracker software was used to track EB3-labeled MT plus ends in order to measure MT growth speeds and MT growth lifetimes in time-lapse images. This methodology allows for the study of MT dynamics and the identification of how localized regulation of MT dynamics within sub-cellular regions contributes to the angiogenic processes of EC branching and migration.

Protocols for Human Umbilical Vein Endothelial Cell (HUVEC) culture, transient transfection of fluorescently-labeled markers of microtubule growth, live-cell imaging and automated analysis of interphase microtubule growth dynamics are detailed.

The physiological process by which new vasculature forms from existing vasculature requires specific signaling events that trigger morphological changes ...