This research was supported by the Intramural Research Program of the NIH, National Institute of Dental and Craniofacial Research.

Part 1: Microscope and Preparation of the Imaging Setup
<ol>
	<li>Any inverted confocal microscope should be suitable for IVM. In this study an Olympus IX81 inverted microscope, equipped with a Fluoview-1000 laser scanning confocal unit was used. To achieve the best possible resolution, the use of high NA objectives, such as the water-immersion UPLSAPO 60x/1.20 NA, and the oil immersion Plan-Apo 60x/1.42 NA is recommended. Oil immersion objectives have higher collection efficiencies and although they require the use of a coverslip between the tissue and the objective, they provide a better imaging quality when used to image areas within 15-20 &#956;m from the surface. Additionally, the use of oil eliminates the concern for water evaporation during longer imaging sessions. In order to maintain the temperature and humidity, the microscope should be fully enclosed and supplied with warm and humidified air. Alternatively, the animal, the microscope stage, and the objective must be warmed by other means. For example, chemical heat pads can be used for the animal and the stage, and specifically designed objective heaters can be used for the objective (Figure 1A). The objective heater should be switched on 30 min before starting the procedure. Note: when the heat pads are use to keep the animal warm a gauze has to place in between the pad and the skin. The skin has to be monitored for burns.</li>
	<li>The standard stages in most of the commercially available confocal microscopes have an opening that is used to insert various holders that are used to fit slides, Petri dishes, and plates of different sizes. In order to perform IVM, a standard stage insert for 35 mm Petri dishes is inverted upside down, covered with a 40 mm glass slide, which is then secured by high vacuum silicone grease or masking tape (Figure 1A). A custom insert can also be manufactured from 3 mm thick aluminum plate. The glass surface is then cleaned with 70% ethanol and swiped with a Kimwipe.</li>
	<li>Manufacture a customized holder for the SGs immobilization (stabilizer). The purpose of the holder is to stabilize the SGs in place and rigidly couple them to the stage and the coverslip. The holder can be made from a 60 mm plastic Petri dish. First, the bottom of the dish is broken away from the walls and cut into rectangles of about 15 x 10 mm. Then, the edges are rounded and polished by using fine sandpaper. Four spacers (1.5-2 mm) are built by removing the rubber tips of 1 ml syringe plungers. By using fine sandpaper, the surface of the spacers and the four corners of the plastic piece are polished. The spacers are then glued by using gelatinous superglue, and later smoothed using fine sandpaper on a flat surface. Note, that for older animals, the size of the holders and the spacers may have to be increased to accommodate slightly larger glands.</li>
	<li>Preparation of the optical coupling gel. 0.3% Carbomer-940 (Table 2) gel is prepared by warming up 100 ml of ultra-pure water and dissolving 5.47 g of D-Sorbitol. Then, 0.3 g of Carbomer-940 are added. The beaker should be covered and placed on a stirring hot plate (lowest heat setting) for several hours or until all the carbomer clumps are completely dissolved. Lastly, Trietanolamine is added drop-wise while gently stirring, until a pH of about 7.4 is reached. The gel is stored at 4 &#176;C 13.</li>
</ol>
Part 2: Animals and Anesthesia
<ol>
	<li>Before conducting any experiments with animals, the protocols should be approved by your local animal care and ethics committee. Importantly, researchers should be properly trained to perform surgical procedures and should consult with their local animal care committee or institution veterinarian before proceeding with the procedure.</li>
	<li>In this study, a mouse model (GFP/mTomato-mouse) derived by crossing FVB mice expressing soluble GFP (GFP-mouse) with C57B6 mice expressing a membrane-targeted peptide fused with the fluorescent protein tandem-Tomato (mTomato-mouse) 6 was used. Note: it usually take 20-30 min for the animal to become fully anesthetized and prepped for imaging</li>
	<li>Mice are housed in a controlled environment in the animal facility and are allowed to acclimate for one week prior to the experiment. This step is necessary for imaging the SGs since any distress results in enhanced secretory activity12. Water and food are provided ad libitum. Males from 2 -5 months old are used for imaging experiments (20-30 g).</li>
	<li>Prior to anesthesia, weigh the animals to determine the proper dose of anesthetic.</li>
	<li>Since the stress induced by administration of injectable anesthetics may impair the secretory activity of the SGs, induce pre-anesthesia by isoflurane inhalation, followed by the injection of anesthetics. To this aim, place the mice very gently into the vaporizer chamber, and turn on the oxygen flow (800-900 mmHg). Set the isoflurane levels to 3% to start the pre-anesthesia induction.</li>
	<li>Once the animals are unconscious, inject a mixture of 100 mg/kg ketamine and 20 mg/kg xylazine into the peritoneal cavity (25 G needle). Anesthetics are freshly prepared by diluting 100 mg/ml xylazine to 20 mg/ml in saline and then mixing it with the ketamine. This mixture is further diluted 1:1 with saline and an appropriate amount is injected based on the animal weight. This dilution helps in preventing accidental overdose. Note that the mixture loses its effectiveness after 20 min.</li>
	<li>Observe the animals for signs of wakefulness and inject more anesthetic mixture if they are coming out of anesthesia (approximately 75% of the initial dose within 45 min after the first injection, and 50% every hour thereafter). Check the depth of the anesthesia every 15 min by pinching the paw and observing the response. Note: Always administer ophthalmic ointment to prevent eye dryness.</li>
	<li>Anesthetics induce hypothermia, which can affect the reproducibility of the experiments and the health of the animal. Therefore, mice should be kept under the heat lamp or on a heated pad for the entire duration of the experiment. Note: the heat lamp should be at least 12-18 inches from the animal to prevent burning</li>
	<li>The body temperature of the animals has to be monitored with a thermometer equipped with a rectal temperature probe.</li>
</ol>
Part 3: Animal Surgery and Positioning for Intravital Microscopy
<ol>
	<li>Prepare the workspace with all the surgical instruments (Table 1) that should be clean and sterile.</li>
	<li>Shave the neck area of the mouse with an electric clipper. Wipe down the ventral side of the neck area of the anesthetized mouse with a gauze sponge soaked in 70% ethanol. Dry the excess with a Kimwipe.</li>
	<li>With dull curved tweezers lift up a small piece of skin at the midline (approximately at the lower end of the masseter muscles), and make a small incision (Figure 1B).</li>
	<li>Insert the scissors into the opening and separate the skin away from the underlying tissue by opening the scissor blades. Avoid damaging the underlying glandular tissue by pointing the scissors up towards the underside of the skin. Repeat this step several times until the skin is separated from the deeper tissues (Figure 1B).</li>
	<li>Using scissors, cut out a strip of loosened skin about 3 mm wide and 10 mm long. The skin should be cut so that the opening starts near the base of the SGs where the nerve, the duct and the blood vessels connect the gland to the rest of the body. After cleaning the wound area, the resultant opening will be oval in shape and measure roughly 8 x 12 mm. Both SGs will be visible after the opening (Figure 1B).</li>
	<li>With a syringe apply the optical coupling gel into the middle of the cut and wipe it towards the outside with sterile gauze. Clean pieces of gauze should be used for each wipe to prevent introduction of hair or blood into the exposed tissue. Repeat this step until all blood and loose hair is removed.</li>
	<li>Using #7 tweezers, separate the connective tissue away from right submandibular gland all the way to the base of the gland (alternatively this protocol can be used on the left SGs). First, find the tip of the SG and grab the connective tissue a few millimeters above the tip of the gland. Tear the connective tissue around the gland without injuring the parenchyma. (Figure 1B). Once the gland is exposed, gently separate the connective tissue from the gland. If bleeding occurs, wash the blood away with saline. Make sure that all the connective tissues are separated and the gland is fully exposed. Keep the gland moist by applying optical coupling gel regularly.</li>
	<li>Prior to bringing the animal to the microscope, place a thick piece of gauze and the heat pack onto the microscope stage to maintain a temperature of 38 &#177; 2 &#176;C. The temperature of the chemical heat pack can be adjusted by the size of the cuts that expose the inner pouch to the atmosphere. Gauze provides some insulation of the heat pack from the metal stage. Cover the top of the heat pack with a thin piece of gauze as well.</li>
	<li>Place the animal on its side (right side for the right gland) and onto the gauze. Position the animal with the submandibular salivary gland placed in the middle of the coverslip (Figure 1C).</li>
	<li>The animal should be secured and stabilized in this position before immobilizing the gland. Place a piece of masking tape on the ventral side of the front right paw of the mouse. Hook the incisors of the mouse by a silk suture (Figure 1C, inset). Create some tension between the silk thread and the paw and attach them onto the stage by masking tape. The SG should be resting naturally in the middle of the coverslip. The head and the thorax should be additionally stabilized by plastic wedges that can be coupled to the stage.</li>
	<li>Place a small piece of lens cleaning tissue over the gland (Figure 1C). Extend the gland slightly and place a customized holder over the gland. Tightly couple the stabilizer to the stage with masking tape. Coupling can be improved by a metal clamp connected to the stage (Figure 1C). Tension and sharp angles should be avoided as they may compromise the blood flow. Immediately after the stabilization inspect the blood flow either by examining the gland visually (it should retain pink coloration) or by epifluorescence. In the GFP/m-tomato mouse, gross alterations of the blood flow can be easily evaluated since both the blood vessels and several of the blood cells are labeled with the m-Tomato.</li>
	<li>Cover the animal with gauze and a heated pad. Measure the temperature of the exposed gland by using a thermometer equipped with a small flexible thermocouple probe placed in contact with one side of the exposed gland.</li>
</ol>
Part 4: Imaging Parameters
<ol>
	<li>Using epifluorescence illumination focus on the surface of the gland by either using the GFP or the RFP filter set. Find the surface of the SG, and assess the blood flow in the capillaries and the overall morphology of the tissue. Signs of tissue damage include membrane blebbing or the appearance of large vacuoles.</li>
	<li>Assess the stability of the tissue either via epifluorescence or by pre-scanning the selected area. A stable preparation is essential, since most of the motion artifacts cannot be removed or corrected for in post-processing data analysis. If the preparation is not stable, adjustments are required.</li>
	<li>To find the proper focal plane image the glands in pre-scanning mode (1X zoom), and gradually move the objective towards the coverslip until the acinar structures, the secretory granules and the plasma membrane become clearly visible (Figure 2).</li>
	<li>The imaging parameters for the time-lapse depend on the individual experiment. To image the dynamics of the secretory granules use a scan speed of 0.5-2 sec/frame, and 256 x 256 pixels with a 0.2 &#956;m/pixel spatial scale (field of view of approximately 50 &#956;m x 50 &#956;m). In order to optimally visualize the granules and the plasma membrane, the optical thickness should be set between 0.9-1.2 &#956;m. To excite the GFP and the tandem-Tomato, one may use a wavelength of 488 nm and 561 nm, respectively. The peak power is set around 0.5 mW for the 488 nm and 1 mW for the 561 nm. This ensures minimal photobleaching.</li>
	<li>The detector settings have to be adjusted to maintain the signal within the dynamic range and depend on the brightness of the specimen.</li>
	<li>Once all the parameters are set, imaging can be started. First, take a snapshot of the field of view to be used as a reference point. While imaging in time lapse, drifting of the tissue in XY and Z may occur. This can be manually compensated by adjusting the scan area and the Z position. The adjustments should be made gradually and by small increments to avoid artifacts. Moreover, the reference image can be used to determine whether photobleaching occurs. If significant photobleaching is detected (more than 15-20% reduction in fluorescence), reduce the laser power by 0.1 mW.</li>
	<li>To stimulate exocytosis, inject subcutaneously 0.1 mg/kg of a freshly prepared solution of isoproterenol in saline. Exocytosis is triggered approximately 1 minute after the injection. Secretory granules fusing with the plasma membrane will be detected by an increase of GFP fluorescence around the granules and the diffusion of the tandem-Tomato peptides into their limiting membranes (Figure 3). We recommend acquiring 3-4 time-lapse sequences in the same area, 10 min each.</li>
	<li>After the imaging session, euthanize the anesthetized animal by CO2 inhalation in a euthanasia chamber (10-12 mmHg for 10 min).</li>
</ol>

<ol>
	<li>Lippincott-Schwartz, J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Emerging+in+vivo+analyses+of+cell+function+using+fluorescence+imaging+(*).">Emerging in vivo analyses of cell function using fluorescence imaging (*).</a> Annu Rev Biochem. 80, 327-332 (2011).</li><li>Cukierman, E., Pankov, R., Stevens, D. R., Yamada, K. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Taking+cell-matrix+adhesions+to+the+third+dimension.">Taking cell-matrix adhesions to the third dimension.</a> Science. 294, 1708-1712 (2001).</li><li>Weigert, R., Sramkova, M., Parente, L., Amornphimoltham, P., Masedunskas, A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Intravital+microscopy:+a+novel+tool+to+study+cell+biology+in+living+animals.">Intravital microscopy: a novel tool to study cell biology in living animals.</a> Histochem Cell Biol. 133, 481-491 (2010).</li><li>Dunn, K. W., 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=Functional+studies+of+the+kidney+of+living+animals+using+multicolor+two-photon+microscopy.">Functional studies of the kidney of living animals using multicolor two-photon microscopy.</a> Am. J. Physiol. Cell Physiol. 283, C905-C916 (2002).</li><li>Masedunskas, A., Porat-Shliom, N., Weigert, R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Regulated+exocytosis:+novel+insights+from+intravital+microscopy.">Regulated exocytosis: novel insights from intravital microscopy.</a> Traffic. 13, 627-634 (2012).</li><li>Masedunskas, A., 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=Role+for+the+actomyosin+complex+in+regulated+exocytosis+revealed+by+intravital+microscopy.">Role for the actomyosin complex in regulated exocytosis revealed by intravital microscopy.</a> Proc. Natl. Acad. Sci. 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So far subcellular structures have been imaged mostly in in vitro (i.e. cell cultures) or in ex vivo (i.e. organ-cultures, tissue slices, acinar preparations) model systems that often do not recapitulate the characteristics of intact live tissues 6. In this respect, the approach presented here represents a major breakthrough since it enables imaging the dynamics of a specific membrane trafficking step (i.e. regulated exocytosis) in living mice.	
	<p class="jove_...

In the past two decades the advent of live microscopy and the use of fluorescent proteins have led to major breakthroughs on every cellular process imaginable, thus advancing our understanding of cell biology 1. This field has benefited tremendously from the use of mammalian cell cultures that are extremely powerful model systems, particularly when it comes to experimental manipulations. However, they do not often provide a true representation of the biology of complex multicellular organisms 2. This issue has begun to be addressed by the development of intravital microscopy (IVM) that has opened the door to investigating key biological questions in fields such as neurobiology, immunology and tumor biology 3. So far, most of the studies based on IVM have been performed at the levels of tissues and individual cells, without providing any information about the dynamics of subcellular compartments. Recently, our laboratory and others have developed IVM techniques capable of imaging subcellular structures in live rodents 4-7, 13-15 and allowing pharmacological and genetic manipulations in vivo. This approach has been used by us to study membrane trafficking in vivo, and more specifically endocytosis and regulated exocytosis 6,7.
Our experimental model system is based on exposing, stabilizing and imaging the submandibular salivary glands (SGs) of anesthetized rodents. The choice of the SGs as a model organ for IVM is due to the fact that the glands are easily accessible by performing a minor surgery, can be externalized without compromising their physiology, and stabilized to reduce the motion artifacts due to heartbeat and respiration. In addition, SGs can be selectively manipulated genetically by injecting either viral or non-viral based vectors through the salivary duct 8,9. Finally, SGs are exocrine glands composed of polarized epithelial cells, which form the acini and the ducts, myoepithelial cells, and a complex population of stromal cells. For this reason, they are an excellent model to study exocytosis, endocytosis, gene delivery, and actin cytoskeleton, as highlighted in our recent studies 10, and offer the opportunity to study aspects of cell biology such as cell polarity, cell division, cell-cell junctions, and ion channels.
In this paper we describe in detail an imaging protocol for achieving subcellular resolution in the epithelium of the SGs of a live mouse. Specifically, we show how to image the secretory granules in the acinar cells of the SGs during regulated exocytosis. As previously shown, upon stimulation with agonists of the beta-adrenergic receptor, the secretory granules fuse with the apical plasma membrane and gradually collapse, releasing their content into the acinar canaliculi 6. Our goal is to provide the basic tools to investigators with minimal experience in surgical procedures and animal handling, so that they can successfully perform IVM at a subcellular resolution. Since the most challenging part in IVM is the preparation of the animal, here we focus on the description of the basic surgical procedures that are utilized to expose and immobilize the SGs without compromising their function. As for the procedures to label subcellular structures, several strategies, such as systemic delivery of fluorescent probes, use of transgenic animals, or a combination of both, have been described elsewhere 7,11.

<table border="1" fo:keep-together.within-page="always">
		<tbody>
		 <tr>
			<td></td>
			<td></td>
			<td></td>
			<td>Reagent</td>
		 </tr>
		 <tr>
			<td>Isoflourane (Forane)</td>
			<td>Baxter</td>
			<td>101936-40</td>
			<td>Handle under chemical hood</td>
		 </tr>
		 <tr>
			<td>Ketamine (ketaved)</td>
			<td>Fort Dodge Animal Health</td>
			<td>57457-034-10</td>
			<td>Stock solution 100 mg/ml</td>
		 </tr>
		 <tr>
			<td>Xylazine (Anased)</td>
			<td>Akorn, Decatur</td>
			<td>61311-481-10</td>
			<td>Stock solution 100 mg/ml</td>
		 </tr>
		 <tr>
			<td>Neomycin/Polymyxin B</td>
			<td>Bausch and Lomb</td>
			<td>24208-785-55</td>
			<td>Use to lubricate the eye of the mouse</td>
		 </tr>
		 <tr>
			<td>Carbomer-940</td>
			<td>Ashalnd, Inc.</td>
			<td>4607-1</td>
			<td></td>
		 </tr>
		 <tr>
			<td>D-Sorbitol</td>
			<td>Sigma-Aldrich</td>
			<td>S1876</td>
			<td></td>
		 </tr>
		 <tr>
			<td>Triethanolamine</td>
			<td>Sigma-Aldrich</td>
			<td>90279</td>
			<td>Add drop wise to prevent the solution to solidify</td>
		 </tr>
		 <tr>
			<td>Isoproternol</td>
			<td>Sigma-Aldrich</td>
			<td>16504</td>
			<td>Prepare fresh solutions in saline when needed. Stocks can be stored at -20 &deg;C</td>
		 </tr>
		 <tr>
			<td></td>
			<td></td>
			<td></td>
			<td>Instrument</td>
		 </tr>
		 <tr>
			<td>Isoflurane V 1.9 (Vaporizer)</td>
			<td>Braintree Scientific</td>
			<td>190AF</td>
			<td></td>
		 </tr>
		 <tr>
			<td>Portable Downdraft table equipped with HEPA filter</td>
			<td>Hazard Technology</td>
			<td>PDDT</td>
			<td></td>
		 </tr>
		 <tr>
			<td>Heat lamp, Model HL1</td>
			<td>Braintree Scientific</td>
			<td>HL-1 US</td>
			<td></td>
		 </tr>
		 <tr>
			<td>MicroTherma 2T Thermometer</td>
			<td>Braintree Scientific</td>
			<td>TW2</td>
			<td></td>
		 </tr>
		 <tr>
			<td>Operating Scissors (11.5 cm straight</td>
			<td>World Precision Instruments </td>
			<td>5003708-12</td>
			<td></td>
		 </tr>
		 <tr>
			<td>#7 curved tip tweezers</td>
			<td>World Precision Instruments</td>
			<td>14187</td>
			<td></td>
		 </tr>
		 <tr>
			<td>Microscissors</td>
			<td>World Precision Instruments</td>
			<td>503365</td>
			<td></td>
		 </tr>
		 <tr>
			<td>Black Braided Silk suture #4.0</td>
			<td>George &amp; Tiemann &amp; Co</td>
			<td>160-1219-4</td>
			<td></td>
		 </tr>
		 <tr>
			<td>Gauze sponges 2" x 2"</td>
			<td>Tyco Healthcare</td>
			<td>9022</td>
			<td></td>
		 </tr>
		 <tr>
			<td>Lens cleaning tissue</td>
			<td>Olympus</td>
			<td>CL-TISSUE (M97) AX6476</td>
			<td></td>
		 </tr>
		</tbody>
 </table>

intravital microscopy for imaging subcellular structures in live mice expressing fluorescent proteins

Here we describe a procedure to image subcellular structures in live rodents that is based on the use of confocal intravital microscopy. As a model organ, we use the salivary glands of live mice since they provide several advantages. First, they can be easily exposed to enable access to the optics, and stabilized to facilitate the reduction of the motion artifacts due to heartbeat and respiration. This significantly facilitates imaging and tracking small subcellular structures. Second, most of the cell populations of the salivary glands are accessible from the surface of the organ. This permits the use of confocal microscopy that has a higher spatial resolution than other techniques that have been used for in vivo imaging, such as two-photon microscopy. Finally, salivary glands can be easily manipulated pharmacologically and genetically, thus providing a robust system to investigate biological processes at a molecular level.
In this study we focus on a protocol designed to follow the kinetics of the exocytosis of secretory granules in acinar cells and the dynamics of the apical plasma membrane where the secretory granules fuse upon stimulation of the beta-adrenergic receptors. Specifically, we used a transgenic mouse that co-expresses cytosolic GFP and a membrane-targeted peptide fused with the fluorescent protein tandem-Tomato. However, the procedures that we used to stabilize and image the salivary glands can be extended to other mouse models and coupled to other approaches to label in vivo cellular components, enabling the visualization of various subcellular structures, such as endosomes, lysosomes, mitochondria, and the actin cytoskeleton.

In the GFP/mTomato mouse, the acini appear as clearly distinct structures, which express cytosolic GFP and membrane-targeted tandem-Tomato peptide (Figure 2, broken line). In individual acini, acinar cells are delineated by the tandem-Tomato peptide. GFP is also detected in the nuclei that are clearly visible inside the acinar cells (Figure 2, arrows). Cytosolic GFP is excluded from the secretory granules that appear as dark circular vesicles of approximately 1-1.5 &mu;m in diameter (<st...

Watch this Scientific Journal Video about Intravital Microscopy for Imaging Subcellular Structures in Live Mice Expressing Fluorescent Proteins at JoVE.com

Intravital Microscopy for Imaging Subcellular Structures in Live Mice Expressing Fluorescent Proteins

Intravital microscopy is a powerful tool that enables imaging various biological processes in live animals. In this article, we present a detailed method for imaging the dynamics of subcellular structures, such as the secretory granules, in the salivary glands of live mice.

Here we describe a procedure to image subcellular structures in live rodents that is based on the use of confocal intravital microscopy. As a model organ, ...

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JoVE Journal

Biology

23.6K vistas.  National Institutes of Health.  Microscopía intravital es una poderosa herramienta que permite obtener imágenes diferentes procesos biológicos en los animales vivos. En este artículo, presentamos un método detallado para obtener imágenes de la dinámica de las estructuras subcelulares, tales como los gránulos de secreción, en las glándulas salivales de ratones vivos. 

Video: Intravital Microscopy for Imaging Subcellular Structures in Live Mice Expressing Fluorescent Proteins

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 ...

Fluorescent Labeling of Drosophila Heart Structures

The Drosophila melanogaster dorsal vessel, or heart, is a tubular structure comprised of a single layer of contractile cardiomyocytes, pericardial cells that align along each side of the heart wall, supportive alary muscles and, in adults, a layer of ventral longitudinal muscle cells. The contractile fibers house conserved constituents of the muscle cytoarchitecture including densely packed bundles of myofibrils and cytoskeletal/submembranous protein complexes, which interact with homologous components of the extracellular matrix. Here we describe a protocol for the fixation and the fluorescent labeling of particular myocardial elements from the hearts of dissected larvae and semi-intact adult Drosophila. Specifically, we demonstrate the labeling of sarcomeric F-actin and of &alpha;-actinin in larval hearts. Additionally, we perform labeling of F-actin and &alpha;-actinin in myosin-GFP expressing adult flies and of &alpha;-actinin and pericardin, a type IV extracellular matrix collagen, in wild type adult hearts. Particular attention is given to a mounting strategy for semi-intact adult hearts that minimizes handling and optimizes the opportunity for maintaining the integrity of the cardiac tubes and the associated tissues. These preparations are suitable for imaging via fluorescent and confocal microscopy. Overall, this procedure allows for careful and detailed analysis of the structural characteristics of the heart from a powerful genetically tractable model system.

Fluorescent Labeling of Drosophila Heart Structures

Here we describe a basic protocol for fluorescent labeling of different elements of heart tubes from larva and adult Drosophila melanogaster. These specimens are well-suited for imaging via fluorescent or confocal microscopy. This technique permits detailed structural analysis of the features of the hearts from a powerful model organism.

The Drosophila melanogaster dorsal vessel, or heart, is a tubular structure comprised of a single layer of contractile cardiomyocytes, pericardial cells ...

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 ...

Transplantation of GFP-expressing Blastomeres for Live Imaging of Retinal and Brain Development in Chimeric Zebrafish Embryos

Cells change extensively in their locations and property during embryogenesis. These changes are regulated by the interactions between the cells and their environment. Chimeric embryos, which are composed of cells of different genetic background, are great tools to study the cell-cell interactions mediated by genes of interest. The embryonic transparency of zebrafish at early developmental stages permits direct visualization of the morphogenesis of tissues and organs at the cellular level. Here, we demonstrate a protocol to generate chimeric retinas and brains in zebrafish embryos and to perform live imaging of the donor cells. The protocol covers the preparation of transplantation needles, the transplantation of GFP-expressing donor blastomeres to GFP-negative hosts, and the examination of donor cell behavior under live confocal microscopy. With slight modifications, this protocol can also be used to study the embryonic development of other tissues and organs in zebrafish. The advantages of using GFP to label donor cells are also discussed. 

We demonstrate a protocol to generate chimeric zebrafish embryos for live imaging cellular behavior during embryogenesis.

Cells change extensively in their locations and property during embryogenesis. These changes are regulated by the interactions between the cells and their ...

Photoconversion of Purified Fluorescent Proteins and Dual-probe Optical Highlighting in Live Cells

Photoconvertible fluorescent proteins (pc-FPs) are a class of fluorescent proteins with "optical highlighter" capability, meaning that the color of fluorescence can be changed by exposure to light of a specific wavelength. Optical highlighting allows noninvasive marking of a subpopulation of fluorescent molecules, and is therefore ideal for tracking single cells or organelles.
Critical parameters for efficient photoconversion are the intensity and the exposure time of the photoconversion light. If the intensity is too low, photoconversion will be slow or not occur at all. On the other hand, too much intensity or too long exposure can photobleach the protein and thereby reduce the efficiency of photoconversion.
This protocol describes a general approach how to set up a confocal laser scanning microscope for pc-FP photoconversion applications. First, we describe a procedure for preparing purified protein droplet samples. This sample format is very convenient for studying the photophysical behavior of fluorescent proteins under the microscope. Second, we will use the protein droplet sample to show how to configure the microscope for photoconversion. And finally, we will show how to perform optical highlighting in live cells, including dual-probe optical highlighting with mOrange2 and Dronpa.

This protocol describes a general approach to perform photoconversion of fluorescent proteins on a confocal laser scanning microscope. We describe procedures for the photoconversion of puried protein samples, as well as for dual-probe optical highlighting in live cells with mOrange2 and Dronpa.

Photoconvertible fluorescent proteins (pc-FPs) are a class of fluorescent proteins with "optical highlighter" capability, meaning that the color of ...

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 ...

In vivo Clonal Tracking of Hematopoietic Stem and Progenitor Cells Marked by Five Fluorescent Proteins using Confocal and Multiphoton Microscopy

We developed and validated a fluorescent marking methodology for clonal tracking of hematopoietic stem and progenitor cells (HSPCs) with high spatial and temporal resolution to study in vivo hematopoiesis using the murine bone marrow transplant experimental model. Genetic combinatorial marking using lentiviral vectors encoding fluorescent proteins (FPs) enabled cell fate mapping through advanced microscopy imaging. Vectors encoding five different FPs: Cerulean, EGFP, Venus, tdTomato, and mCherry were used to concurrently transduce HSPCs, creating a diverse palette of color marked cells. Imaging using confocal/two-photon hybrid microscopy enables simultaneous high resolution assessment of uniquely marked cells and their progeny in conjunction with structural components of the tissues. Volumetric analyses over large areas reveal that spectrally coded HSPC-derived cells can be detected non-invasively in various intact tissues, including the bone marrow (BM), for extensive periods of time following transplantation. Live studies combining video-rate multiphoton and confocal time-lapse imaging in 4D demonstrate the possibility of dynamic cellular and clonal tracking in a quantitative manner.

In vivo Clonal Tracking of Hematopoietic Stem and Progenitor Cells Marked by Five Fluorescent Proteins using Confocal and Multiphoton Microscopy

Combinatorial 5 fluorescent proteins marking of hematopoietic stem and progenitor cells allows in vivo clonal tracking via confocal and two-photon microscopy, providing insights into bone marrow hematopoietic architecture during regeneration. This method allows non-invasive fate mapping of spectrally-coded HSPCs-derived cells in intact tissues for extensive periods of time following transplantation.

We developed and validated a fluorescent marking methodology for clonal tracking of hematopoietic stem and progenitor cells (HSPCs) with high spatial and ...

Simultaneous Measurement of Mitochondrial Calcium and Mitochondrial Membrane Potential in Live Cells by Fluorescent Microscopy

Apart from their essential role in generating ATP, mitochondria also act as local calcium (Ca2+) buffers to tightly regulate intracellular Ca2+ concentration. To do this, mitochondria utilize the electrochemical potential across their inner membrane (&#916;&#936;m) to sequester Ca2+. The influx of Ca2+ into the mitochondria stimulates three rate-limiting dehydrogenases of the citric acid cycle, increasing electron transfer through the oxidative phosphorylation (OXPHOS) complexes. This stimulation maintains &#916;&#936;m, which is temporarily dissipated as the positive calcium ions cross the mitochondrial inner membrane into the mitochondrial matrix.
We describe here a method for simultaneously measuring mitochondria Ca2+ uptake and &#916;&#936;m in live cells using confocal microscopy. By permeabilizing the cells, mitochondrial Ca2+ can be measured using the fluorescent Ca2+ indicator Fluo-4, AM, with measurement of &#916;&#936;m using the fluorescent dye tetramethylrhodamine, methyl ester, perchlorate (TMRM). The benefit of this system is that there is very little spectral overlap between the fluorescent dyes, allowing accurate measurement of mitochondrial Ca2+ and &#916;&#936;m simultaneously. Using the sequential addition of Ca2+ aliquots, mitochondrial Ca2+ uptake can be monitored, and the concentration at which Ca2+ induces mitochondrial membrane permeability transition and the loss of &#916;&#936;m determined.

Mitochondria can utilize the electrochemical potential across their inner membrane (&#916;&#936;m) to sequester calcium (Ca2+), allowing them to shape cytosolic Ca2+ signaling within the cell. We describe a method for simultaneously measuring mitochondria Ca2+ uptake and &#916;&#936;m in live cells using fluorescent dyes and confocal microscopy.

Apart from their essential role in generating ATP, mitochondria also act as local calcium (Ca2+) buffers to tightly regulate intracellular Ca2+ ...

Fluorescence Recovery after Photobleaching of Yellow Fluorescent Protein Tagged p62 in Aggresome-like Induced Structures

Fluorescence recovery after photobleaching (FRAP)&#160;is a microscopy technique that can be used to quantify protein mobility in live cells. In a typical FRAP experiment, steady-state fluorescence is observed by repeated imaging with low-intensity laser light. Subsequently, the fluorescent molecules are rapidly and irreversibly impaired via brief exposure to high-intensity laser light. Information about protein mobility is obtained by monitoring the recovery of fluorescence. We used FRAP to determine the mobility of p62 in aggresome-like induced structures (ALIS) in murine macrophages after stimulation with lipopolysaccharide (LPS). Because many existing FRAP protocols are either incomplete or complex, our goal was to provide a comprehensive, practical, and straightforward step-by-step protocol for FRAP experiments with live cells. Here, we describe RAW264.7 macrophage transfection with yellow fluorescent protein-p62 (YFP-p62), induction of ALIS by exposing the cells to LPS, and a step-by-step method for collecting prebleach and postbleach FRAP images and data analysis. Finally, we discuss important factors to consider when conducting a FRAP experiment.

We describe a comprehensive and practical protocol for fluorescence recovery after photobleaching experiments with live cells. Although the protocol was used to measure the mobility of yellow fluorescent protein-tagged p62 in aggresome-like induced structures, it can be applied to a variety of microscopy systems and fluorescent proteins.

Fluorescence recovery after photobleaching (FRAP)&#160;is a microscopy technique that can be used to quantify protein mobility in live cells. In a typical ...

Super-Resolution Live Cell Imaging of Subcellular Structures

There has long been a crucial tradeoff between spatial and temporal resolution in imaging. Imaging beyond the diffraction limit of light has traditionally been restricted to be used only on fixed samples or live cells outside of tissue labeled with strong fluorescent signal. Current super-resolution live cell imaging techniques require the use of special fluorescence probes, high illumination, multiple image acquisitions with post-acquisition processing, or often a combination of these processes. These prerequisites significantly limit the biological samples and contexts that this technique can be applied to.
Here we describe a method to perform super-resolution (~140 nm XY-resolution) time-lapse fluorescence live cell imaging in situ. This technique is also compatible with low fluorescent intensity, for example, EGFP or mCherry endogenously tagged at lowly expressed genes. As a proof-of-principle, we have used this method to visualize multiple subcellular structures in the Drosophila testis. During tissue preparation, both the cellular structure and tissue morphology are maintained within the dissected testis. Here, we use this technique to image microtubule dynamics, the interactions between microtubules and the nuclear membrane, as well as the attachment of microtubules to centromeres.
This technique requires special procedures in sample preparation, sample mounting and immobilizing of specimens. Additionally, the specimens must be maintained for several hours after dissection without compromising cellular function and activity. While we have optimized the conditions for live super-resolution imaging specifically in Drosophila male germline stem cells (GSCs) and progenitor germ cells in dissected testis tissue, this technique is broadly applicable to a variety of different cell types. The ability to observe cells under their physiological conditions without sacrificing either spatial or temporal resolution will serve as an invaluable tool to researchers seeking to address crucial questions in cell biology.

Presented here is a protocol for super-resolution live-cell imaging in intact tissue. We have standardized the conditions for imaging a highly sensitive adult stem cell population in its native tissue environment. This technique involves balancing temporal and spatial resolution to allow for the direct observation of biological phenomena in live tissue.

There has long been a crucial tradeoff between spatial and temporal resolution in imaging. Imaging beyond the diffraction limit of light has traditionally ...