Name: live cell imaging of primary rat neonatal cardiomyocytes following adenoviral and lentiviral transduction using confocal spinning disk microscopy
Uploaded: 2014-06-24T14:00:00
Description: Watch this Scientific Journal Video about Live Cell Imaging of Primary Rat Neonatal Cardiomyocytes Following Adenoviral and Lentiviral Transduction Using Confocal Spinning Disk Microscopy at JoVE.com

We would like to thank Dr. Alengo Nyamay'antu and Dr. Ilse Timmerman for their advices about lentiviral packaging. This work is supported by an American Heart Association Scientist Development Grant 10SDG4170137.

1. Isolation of rat neonatal cardiomyocytes&#160;
<ol>
	<li>Use Dulbecco&#39;s modi&#64257;ed Eagle&#39;s medium (DMEM) and minimum essential medium (MEM) supplemented with fetal bovine serum (FBS) and penicillin/streptomycin (P/S) as the culture medium. Add bromodeoxyuridine (BrdU) to the medium to prevent growth of fibroblasts for the first 2 days after the isolation.</li>
</ol>
<table border="1" cellpadding="0" cellspacing="0">
	<tbody>
		<tr>
			<td></td>
			<td>Base medium</td>
			<td>FBS</td>
			<td>BrdU (mM)</td...

<ol>
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Primary cardiomyocytes isolated from neonatal rats have long been used to study cardiomyocyte functions in vitro. This protocol describes a method for the isolation of neonatal cardiomyocytes from rat pups using a two-step enzyme digestion method, first digesting with trypsin O/N at 4 &#176;C and then purified collagenase. One advantage of employing the purified collagenase step is that the same grade of enzyme is used for all isolations. Thus, the quality and amount of isolated cells is consistent from experime...

Primary rat neonatal cardiomyocytes have long been used for investigating cardiomyocyte function in vitro1. They are easy to isolate from rat pups by several well established methods2-4. The most common method employs collagenase or trypsin to digest connective tissue of the heart prior to cell isolation. Researchers have also developed methods for isolating cardiomyocytes from adult rodents5-8 as well as neonatal mice9,10.
This protocol describes a method for isolating cardiomyocytes from neonatal rat pups, employing a two-step enzyme digestion procedure. Trypsin is first used O/N at 4&#160;&#176;C, and then purified collagenase at 37&#160;&#176;C. Incubation of the heart tissue with trypsin O/N at 4&#160;&#176;C&#160;reduces the steps necessary to harvest cells compared to methods using sequential incubations in a warm enzyme solution2. In addition, by using purified collagenase rather than crude enzymes, lot-to-lot variability can be eliminated, thus providing enhanced reproducibility.
Functional studies of a particular protein often employ a protein expression system using an adenovirus11-13 and/or a lentivirus14-16. [CAUTIONARY NOTE] The viral production and manipulation should be carried out according to the NIH guidelines.
The adenovirus does not integrate into the host genome. It has a very high efficiency of transduction in most cell types, including dividing cells and non-dividing cells, as well as primary cells and established cell lines. This makes the adenovirus a reliable vector for gene expression. High levels of the protein encoded by the adenovirus vector develop within 48 hr following transduction, and they can last for several weeks17. However, one drawback to using an adenovirus for protein expression is that the development of a recombinant adenovirus is both complicated and time consuming. This drawback explains in part why many researchers have been turning to lentiviruses for recombinant gene expression. Unlike adenoviral constructs, generating a lentiviral construct is quick and easy. Although lentiviruses generally have lower efficiencies of transduction than adenoviruses, in both dividing and non-dividing cells, they do integrate into the host genome. Consequently, expression of the transduced gene is more stable for lentiviruses than for adenoviruses.
Due to technological advances in the field of microscopy, it is much easier to capture live cell images of cells expressing recombinant proteins. This holds true even at video rate speeds of acquisition. This allows the investigator to determine how particular alterations in the protein of interest functionally impact the cell in real time. The confocal spinning disk microscope has several key features that make it an optimal technique for live cell imaging18,19. The Yokogawa spinning disc allows for much more rapid image acquisition, while at the same time utilizing far less laser power than point scanning confocal microscopes. Both of these special features are due to the spinning disk, which contains numerous confocal holes through which the laser passes simultaneously to the samples. During acquisition, the disk itself spins quickly and continuously18-20. By using the autofocus system of the microscope, stable focus is maintained over long periods of time. This allows researchers to take well-focused live cell images. Acquired images are played back as a movie file. The images are analyzed using image analysis software such as ImageJ21,22, FIJI23, or other commercially available software.

<table border="0" cellpadding="0" cellspacing="0">
	<tbody>
		<tr>
			<td>1 scissors for decapitation</td>
			<td>WPI</td>
			<td>501749</td>
			<td>Autoclave before use</td>
		</tr>
		<tr>
			<td>1 fine scissors for heart isolation and chopping</td>
			<td>WPI</td>
			<td>14393</td>
			<td>Autoclave before use</td>
		</tr>
		<tr>
			<td>2 fine forceps (Dumont No. 5)</td>
			<td>Sigma</td>
			<td>F6521</td>
			<td>Autoclave before use</td>
		</tr>
		<tr>
			<td>Three sterilized 10 cm plastic dishes</td>
			<td>Sigma</td>
			<td>CLS430165</td>
			<td>for hearts isolation</td>
		</tr>
		<tr>
			<td>3.5 cm glass bottom culture dishes</td>
			<td>MatTek</td>
			<td>P35G-1.5-20-C</td>
			<td>for final plating of cardiomyocytes for future live cell imaging. micro-Dishes from ibidi are an acceptable alternative.</td>
		</tr>
		<tr>
			<td>3.5 cm glass bottom culture dishes</td>
			<td>MatTek</td>
			<td>P35G-0.17-14-C</td>
			<td>for TIRF or high resolution image</td>
		</tr>
		<tr>
			<td>Ethanol solution, 70 % (v/v) in water</td>
			<td>Sigma</td>
			<td>E7148</td>
			<td></td>
		</tr>
		<tr>
			<td>2% gelatin</td>
			<td>Sigma</td>
			<td>G1393</td>
			<td></td>
		</tr>
		<tr>
			<td>One sterilized 10 cm plastic dish</td>
			<td>Sigma</td>
			<td>CLS430165</td>
			<td>for trypsinization</td>
		</tr>
		<tr>
			<td>Aluminium foil</td>
			<td>any brand</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>parafilm</td>
			<td>Sigma</td>
			<td>P7543</td>
			<td></td>
		</tr>
		<tr>
			<td>Two 10 cm plastic cell culture dish</td>
			<td>Sigma</td>
			<td>CLS430165</td>
			<td>for selection</td>
		</tr>
		<tr>
			<td>Auto pipette</td>
			<td>Drummond Scientific&#160;</td>
			<td>4-000-300</td>
			<td>for trituration</td>
		</tr>
		<tr>
			<td>Cell counter</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>&#160; Cellometer, automated cell counter</td>
			<td>nexcelom</td>
			<td></td>
			<td>to check and count cells</td>
		</tr>
		<tr>
			<td>&#160; Microscope and Hematocytometer</td>
			<td>any brand</td>
			<td></td>
			<td>to check and count cells</td>
		</tr>
		<tr>
			<td>Trypan Blue Solution, 0.4%</td>
			<td>invitrogen</td>
			<td>15250-061</td>
			<td></td>
		</tr>
		<tr>
			<td>CO2 incubator</td>
			<td>Sanyo</td>
			<td>MCO-19AIC</td>
			<td></td>
		</tr>
		<tr>
			<td>incubating orbital shaker</td>
			<td>Sigma</td>
			<td>Z673129</td>
			<td>to shake heart tissue with collagenase at 37 C at 170-200 rpm</td>
		</tr>
		<tr>
			<td>10 mg/ml BrdU solution</td>
			<td>BD Pharmingen</td>
			<td>550891</td>
			<td></td>
		</tr>
		<tr>
			<td>DMEM, High Glucose</td>
			<td>invitrogen</td>
			<td>41965039</td>
			<td>Mix medium as indicated in the protocol and warm before use</td>
		</tr>
		<tr>
			<td>MEM</td>
			<td>invitrogen</td>
			<td>31095029</td>
			<td>Mix medium as indicated in the protocol and warm before use</td>
		</tr>
		<tr>
			<td>Fetal Bovine Serum</td>
			<td>invitrogen</td>
			<td>26140079</td>
			<td></td>
		</tr>
		<tr>
			<td>Penicillin-Streptomycin (10,000 U/mL)&#160;</td>
			<td>invitrogen</td>
			<td>15140-122</td>
			<td></td>
		</tr>
		<tr>
			<td></td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Section 2 lentiviral transduction</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>three packaging plasmids</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>&#160;pMDLg/pRRE</td>
			<td>Addgene</td>
			<td>12251</td>
			<td></td>
		</tr>
		<tr>
			<td>&#160;pRSV-Rev</td>
			<td>Addgene</td>
			<td>12253</td>
			<td></td>
		</tr>
		<tr>
			<td>&#160;pMD2.G</td>
			<td>Addgene</td>
			<td>12259</td>
			<td></td>
		</tr>
		<tr>
			<td>The lentiviral transfer vector, pLVX-IRES-Puro</td>
			<td>Clontech</td>
			<td>632183</td>
			<td></td>
		</tr>
		<tr>
			<td>Opti-MEM (serum-free medium)</td>
			<td>invitrogen</td>
			<td>31985070</td>
			<td></td>
		</tr>
		<tr>
			<td>transfection reagent</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>&#160; polyethyleneimine&#8220;Max&#8221;, (Mw 40,000) - High Potency Linear PEI (Equivalent to Mw 25,000 in Free Base Form)&#160;</td>
			<td>Polysciences</td>
			<td>24765-2</td>
			<td>It can be substituted with X-tremeGENE 9 from Roche</td>
		</tr>
		<tr>
			<td>&#160; X-tremeGENE 9</td>
			<td>Roche</td>
			<td>6365779001</td>
			<td>substitute for PEI as transfection reagent</td>
		</tr>
		<tr>
			<td>chloroquine</td>
			<td>Sigma</td>
			<td>C6628</td>
			<td>dissolve in water and make 100 mM stock solution. Inhibition of endosomal acidification can be achieved with 10-100 &#956;M Chloroquine.</td>
		</tr>
		<tr>
			<td>HEPES</td>
			<td>Sigma</td>
			<td>H3375</td>
			<td></td>
		</tr>
		<tr>
			<td>10 ml Luer-Lok syringe, sterilized</td>
			<td>BD</td>
			<td>309604</td>
			<td></td>
		</tr>
		<tr>
			<td>0.45 um filters</td>
			<td>Sigma</td>
			<td>F8677</td>
			<td>use only cellulose acetate or polyethersulfone (PES) (low protein binding) filters. Do not use nitrocellulose filters. Nitrocellulose binds surface proteins on the lentiviral envelope and destroys the virus.</td>
		</tr>
		<tr>
			<td>Hexadimethrine bromide</td>
			<td>Sigma</td>
			<td>H9268</td>
			<td>dissolve in water and make 8mg/ml stock solution, then filter it to sterilize.</td>
		</tr>
		<tr>
			<td>polyethylene glicol 6000</td>
			<td>Sigma</td>
			<td>81260</td>
			<td></td>
		</tr>
		<tr>
			<td>Sodium chloride</td>
			<td>Sigma</td>
			<td>S9888</td>
			<td></td>
		</tr>
		<tr>
			<td>sodium hydroxide</td>
			<td>Sigma</td>
			<td>S5881</td>
			<td></td>
		</tr>
		<tr>
			<td></td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Section 3 Adenoviral transduction</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>HEK 293T cells</td>
			<td>ATCC</td>
			<td>CRL-11268</td>
			<td></td>
		</tr>
		<tr>
			<td>Some 10 cm cell culture dishes</td>
			<td>Sigma</td>
			<td>CLS430165</td>
			<td></td>
		</tr>
		<tr>
			<td>96-Well Microplate with lid, flat-bottom, tissue culture, sterile</td>
			<td>BD Falcon</td>
			<td>353072</td>
			<td>for titration</td>
		</tr>
		<tr>
			<td>Multichannel piptte, 10-100 ul, 8-channel</td>
			<td>eppendorf</td>
			<td>3122 000.035</td>
			<td></td>
		</tr>
		<tr>
			<td></td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Section 4 live cell imaging</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Spinning disk confocal microspopy</td>
			<td>PerkinElmer</td>
			<td>L7267000</td>
			<td></td>
		</tr>
		<tr>
			<td>a temperature controlled chamber</td>
			<td>any brand</td>
			<td></td>
			<td>to keep temperature at 37 C</td>
		</tr>
		<tr>
			<td>a CO2 environmental system</td>
			<td>any brand</td>
			<td></td>
			<td>optional to maintain CO2 concentration optimal</td>
		</tr>
		<tr>
			<td>Medium</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>&#160; CO2 Independent Medium, No Glutamine</td>
			<td>invitrogen</td>
			<td>18045-054&#160;&#160;</td>
			<td>for long time time-lapse imaging</td>
		</tr>
		<tr>
			<td>&#160; DMEM, High Glucose, HEPES, no Phenol Red&#160;</td>
			<td>invitrogen</td>
			<td>21063-029</td>
			<td>for long time time-lapse imaging</td>
		</tr>
	</tbody>
</table>

live cell imaging of primary rat neonatal cardiomyocytes following adenoviral and lentiviral transduction using confocal spinning disk microscopy

Primary rat neonatal cardiomyocytes are useful in basic in vitro cardiovascular research because they can be easily isolated in large numbers in a single procedure. Due to advances in microscope technology it is relatively easy to capture live cell images for the purpose of investigating cellular events in real time with minimal concern regarding phototoxicity to the cells. This protocol describes how to take live cell timelapse images of primary rat neonatal cardiomyocytes using a confocal spinning disk microscope following lentiviral and adenoviral transduction to modulate properties of the cell. The application of two different types of viruses makes it easier to achieve an appropriate transduction rate and expression levels for two different genes. Well focused live cell images can be obtained using the microscope&#8217;s autofocus system, which maintains stable focus for long time periods. Applying this method, the functions of exogenously engineered proteins expressed in cultured primary cells can be analyzed. Additionally, this system can be used to examine the functions of genes through the use of siRNAs as well as of chemical modulators.

To illustrate the technique, a lentivirus encoding EGFP (enhanced green fluorescent protein)-tagged Cx43 (Connexin43) or a mutated Cx43 32 was used to express EGFP-tagged proteins in cells, and an adenovirus encoding FGFR1DN (fibroblast growth factor receptor 1 dominant negative) was used to shut down FGF signaling in the cell 33-35. Three days following the isolation of cardiomyocytes, the isolated cardiomyocytes were transduced with lentivirus in order to express EGFP-tagged proteins in the cells....

Watch this Scientific Journal Video about Live Cell Imaging of Primary Rat Neonatal Cardiomyocytes Following Adenoviral and Lentiviral Transduction Using Confocal Spinning Disk Microscopy at JoVE.com

Live Cell Imaging of Primary Rat Neonatal Cardiomyocytes Following Adenoviral and Lentiviral Transduction Using Confocal Spinning Disk Microscopy

This protocol describes a method of live cell imaging using primary rat neonatal cardiomyocytes following lentiviral and adenoviral transduction using confocal spinning disk microscopy. This enables detailed observations of cellular processes in living cardiomyocytes.

Primary rat neonatal cardiomyocytes are useful in basic in vitro cardiovascular research because they can be easily isolated in large numbers in a single ...

live-cell-imaging-primary-rat-neonatal-cardiomyocytes-following

Research

JoVE Journal

Biology

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

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

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

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

Live Imaging of the Zebrafish Embryonic Brain by Confocal Microscopy

In this video, we demonstrate the method our lab has developed to analyze the cell shape changes and rearrangements required to bend and fold the developing zebrafish brain (Gutzman et al, 2008). Such analysis affords a new understanding of the underlying cell biology required for development of the 3D structure of the vertebrate brain, and significantly increases our ability to study neural tube morphogenesis. The embryonic zebrafish brain is shaped beginning at 18 hours post fertilization (hpf) as the ventricles within the neuroepithelium inflate. By 24 hpf, the initial steps of neural tube morphogenesis are complete. Using the method described here, embryos at the one cell stage are injected with mRNA encoding membrane-targeted green fluorescent protein (memGFP). After injection and incubation, the embryo, now between 18 and 24 hpf, is mounted, inverted, in agarose and imaged by confocal microscopy. Notably, the zebrafish embryo is transparent making it an ideal system for fluorescent imaging. While our analyses have focused on the midbrain-hindbrain boundary and the hindbrain, this method could be extended for analysis of any region in the zebrafish to a depth of 
80-100 &mu;m.

In this video, we demonstrate a method by which to analyze the developing vertebrate brain in live zebrafish embryos at single cell resolution by confocal microscopy. This includes the method by which we inject the single-cell zebrafish embryo and subsequently mount and image the developing brain.

In this video, we demonstrate the method our lab has developed to analyze the cell shape changes and rearrangements required to bend and fold the ...

Fluorescence-based Measurement of Store-operated Calcium Entry in Live Cells: from Cultured Cancer Cell to Skeletal Muscle Fiber

Store operated Ca2+ entry (SOCE), earlier termed capacitative Ca2+ entry, is a tightly regulated mechanism for influx of extracellular Ca2+ into cells to replenish depleted endoplasmic reticulum (ER) or sarcoplasmic reticulum (SR) Ca2+ stores1,2. Since Ca2+ is a ubiquitous second messenger, it is not surprising to see that SOCE plays important roles in a variety of cellular processes, including proliferation, apoptosis, gene transcription and motility. Due to its wide occurrence in nearly all cell types, including epithelial cells and skeletal muscles, this pathway has received great interest3,4. However, the heterogeneity of SOCE characteristics in different cell types and the physiological function are still not clear5-7.
The functional channel properties of SOCE can be revealed by patch-clamp studies, whereas a large body of knowledge about this pathway has been gained by fluorescence-based intracellular Ca2+ measurements because of its convenience and feasibility for high-throughput screening. The objective of this report is to summarize a few fluorescence-based methods to measure the activation of SOCE in monolayer cells, suspended cells and muscle fibers5,8-10. The most commonly used of these fluorescence methods is to directly monitor the dynamics of intracellular Ca2+ using the ratio of F340nm and F380nm (510 nm for emission wavelength) of the ratiometric Ca2+ indicator Fura-2. To isolate the activity of unidirectional SOCE from intracellular Ca2+ release and Ca2+ extrusion, a Mn2+ quenching assay is frequently used. Mn2+ is known to be able to permeate into cells via SOCE while it is impervious to the surface membrane extrusion processes or to ER uptake by Ca2+ pumps due to its very high affinity with Fura-2. As a result, the quenching of Fura-2 fluorescence induced by the entry of extracellular Mn2+ into the cells represents a measurement of activity of SOCE9. Ratiometric measurement and the Mn+2 quenching assays can be performed on a cuvette-based spectrofluorometer in a cell population mode or in a microscope-based system to visualize single cells. The advantage of single cell measurements is that individual cells subjected to gene manipulations can be selected using GFP or RFP reporters, allowing studies in genetically modified or mutated cells. The spatiotemporal characteristics of SOCE in structurally specialized skeletal muscle can be achieved in skinned muscle fibers by simultaneously monitoring the fluorescence of two low affinity Ca2+ indicators targeted to specific compartments of the muscle fiber, such as Fluo-5N in the SR and Rhod-5N in the transverse tubules9,11,12.

The extent of store-operated Ca2+ entry (SOCE) can be monitored using fluorescent Ca2+ indicators. Mn2+ quenching of such indicators assays SOCE in cultured cells and skeletal muscle fibers. A technique allowing spatial and temporal resolution of SOCE by confocal imaging of mechanically skinned muscle fibers is also described.

Store operated Ca2+ entry (SOCE), earlier termed capacitative Ca2+ entry, is a tightly regulated mechanism for influx of extracellular Ca2+ into cells to ...

Isolation and Culture of Neonatal Mouse Cardiomyocytes

Cultured neonatal cardiomyocytes have long been used to study myofibrillogenesis and myofibrillar functions. Cultured cardiomyocytes allow for easy investigation and manipulation of biochemical pathways, and their effect on the biomechanical properties of spontaneously beating cardiomyocytes.	
The following 2-day protocol describes the isolation and culture of neonatal mouse cardiomyocytes. We show how to easily dissect hearts from neonates, dissociate the cardiac tissue and enrich cardiomyocytes from the cardiac cell-population. We discuss the usage of different enzyme mixes for cell-dissociation, and their effects on cell-viability. The isolated cardiomyocytes can be subsequently used for a variety of morphological, electrophysiological, biochemical, cell-biological or biomechanical assays. We optimized the protocol for robustness and reproducibility, by using only commercially available solutions and enzyme mixes that show little lot-to-lot variability. We also address common problems associated with the isolation and culture of cardiomyocytes, and offer a variety of options for the optimization of isolation and culture conditions.

Primary mouse cardiomyocyte cultures are one of the pivotal tools for the investigation of myofibrillar organization and function. The following protocol describes the isolation and culture of primary cardiomyocytes from neonatal mouse hearts. The resulting cardiomyocyte cultures may be subsequently used for a variety of biomechanical, biochemical and cell-biological assays.

Cultured neonatal cardiomyocytes have long been used to study  myofibrillogenesis and myofibrillar functions. Cultured cardiomyocytes allow  for easy ...

Analyzing Craniofacial Morphogenesis in Zebrafish Using 4D Confocal Microscopy

Time-lapse imaging is a technique that allows for the direct observation of the process of morphogenesis, or the generation of shape. Due to their optical clarity and amenability to genetic manipulation, the zebrafish embryo has become a popular model organism with which to perform time-lapse analysis of morphogenesis in living embryos. Confocal imaging of a live zebrafish embryo requires that a tissue of interest is persistently labeled with a fluorescent marker, such as a transgene or injected dye. The process demands that the embryo is anesthetized and held in place in such a way that healthy development proceeds normally. Parameters for imaging must be set to account for three-dimensional growth and to balance the demands of resolving individual cells while getting quick snapshots of development. Our results demonstrate the ability to perform long-term in vivo imaging of fluorescence-labeled zebrafish embryos and to detect varied tissue behaviors in the cranial neural crest that cause craniofacial abnormalities. Developmental delays caused by anesthesia and mounting are minimal, and embryos are unharmed by the process. Time-lapse imaged embryos can be returned to liquid medium and subsequently imaged or fixed at later points in development. With an increasing abundance of transgenic zebrafish lines and well-characterized fate mapping and transplantation techniques, imaging any desired tissue is possible. As such, time-lapse in vivo imaging combines powerfully with zebrafish genetic methods, including analyses of mutant and microinjected embryos.

Time-lapse confocal imaging is a powerful technique useful for characterizing embryonic development. Here, we describe the methodology and characterize craniofacial morphogenesis in wild-type, as well as pdgfra, smad5, and smo mutant embryos.

Time-lapse imaging is a technique that allows for the direct observation of the process of morphogenesis, or the generation of shape. Due to their optical ...

Isolation, Culture and Transduction of Adult Mouse Cardiomyocytes

Cultured cardiomyocytes can be used to study cardiomyocyte biology using techniques that are complementary to in vivo systems. For example, the purity and accessibility of in vitro culture enables fine control over biochemical analyses, live imaging, and electrophysiology. Long-term culture of cardiomyocytes offers access to additional experimental approaches that cannot be completed in short term cultures. For example, the in vitro investigation of dedifferentiation, cell cycle re-entry, and cell division has thus far largely been restricted to rat cardiomyocytes, which appear to be more robust in long-term culture. However, the availability of a rich toolset of transgenic mouse lines and well-developed disease models make mouse systems attractive for cardiac research. Although several reports exist of adult mouse cardiomyocyte isolation, few studies demonstrate their long-term culture. Presented here, is a step-by-step method for the isolation and long-term culture of adult mouse cardiomyocytes. First, retrograde Langendorff perfusion is used to efficiently digest the heart with proteases, followed by gravity sedimentation purification. After a period of dedifferentiation following isolation, the cells gradually attach to the culture and can be cultured for weeks. Adenovirus cell lysate is used to efficiently transduce the isolated cardiomyocytes. These methods provide a simple, yet powerful model system to study cardiac biology.

This protocol describes a step-by-step method for the reproducible isolation and long-term culture of adult mouse cardiomyocytes with high yield, purity, and viability.

Cultured cardiomyocytes can be used to study cardiomyocyte biology using techniques that are complementary to in vivo systems. For example, the purity and ...

Live Cell Imaging and 3D Analysis of Angiotensin Receptor Type 1a Trafficking in Transfected Human Embryonic Kidney Cells Using Confocal Microscopy

Live-cell imaging is used to simultaneously capture time-lapse images of angiotensin type 1a receptors (AT1aR) and intracellular compartments in transfected human embryonic kidney-293 (HEK) cells following stimulation with angiotensin II (Ang II). HEK cells are transiently transfected with plasmid DNA containing AT1aR tagged with enhanced green fluorescent protein (EGFP). Lysosomes are identified with a red fluorescent dye. Live-cell images are captured on a laser scanning confocal microscope after Ang II stimulation and analyzed by software in three dimensions (3D, voxels) over time. Live-cell imaging enables investigations into receptor trafficking and avoids confounds associated with fixation, and in particular, the loss or artefactual displacement of EGFP-tagged membrane receptors. Thus, as individual cells are tracked through time, the subcellular localization of receptors can be imaged and measured. Images must be acquired sufficiently rapidly to capture rapid vesicle movement. Yet, at faster imaging speeds, the number of photons collected is reduced. Compromises must also be made in the selection of imaging parameters like voxel size in order to gain imaging speed. Significant applications of live-cell imaging are to study protein trafficking, migration, proliferation, cell cycle, apoptosis, autophagy and protein-protein interaction and dynamics, to name but a few.

Here we present a protocol to image cells expressing green fluorescent protein-tagged angiotensin type 1a receptors during endocytosis initiated by angiotensin II treatment. This technique includes labeling lysosomes with a second fluorescent marker, and then utilizing software to analyze the co-localization of receptor and lysosome in three dimensions over time.

Live-cell imaging is used to simultaneously capture time-lapse images of angiotensin type 1a receptors (AT1aR) and intracellular compartments in ...

Measurement of Microtubule Dynamics by Spinning Disk Microscopy in Monopolar Mitotic Spindles

We describe a modification of an established method to determine microtubule dynamics in living cells. The protocol is based on the expression of a genetically encoded marker for the positive ends of microtubules (EB3 labelled with tdTomato fluorescent protein) and high-speed, high-resolution, live-cell imaging using spinning disk confocal microscopy. Cell cycle synchronization and increased density of microtubules are achieved by inhibiting centrosomal separation in mitotic cells, and analysis of growth is performed using open-source U-Track software. The use of a bright and red-shifted fluorescent protein, in combination with the lower laser power and reduced exposure time required for spinning disk microscopy reduce phototoxicity and the probability of light-induced artifacts. This allows for imaging a larger number of cells in the same preparation while maintaining the cells in a growth medium under standard culture conditions. Because the analysis is performed in a supervised automatic fashion, the results are statistically robust and reproducible.

Here we present a robust and detailed method of microtubule dynamics analysis in cells synchronized in prometaphase using live-cell spinning disk confocal microscopy and MATLAB-based image processing.

We describe a modification of an established method to determine microtubule dynamics in living cells. The protocol is based on the expression of a ...

Autofluorescence Imaging to Evaluate Red Algae Physiology

Red algae (Rhodophyta) contain phycobiliproteins and colonize habitats with dim light, however some (e.g., some Chroothece species) can also develop in full sunshine. Most rhodophytes are red, however some can appear bluish, depending on the proportion of blue and red biliproteins (phycocyanin and phycoerythrin). Different phycobiliproteins can capture light at diverse wavelengths and transmit it to chlorophyll a, which makes photosynthesis under very different light conditions possible. These pigments respond to habitat changes in light, and their autofluorescence can help to study biological processes. Using Chroothece mobilis as a model organism and the spectral lambda scan mode in a confocal microscope, the adaptation of photosynthetic pigments to different monochromatic lights was studied at the cellular level to guess the species' optimal growth conditions. The results showed that, even when the studied strain was isolated from a cave, it adapted to both dim and medium light intensities. The presented method is especially useful for studying photosynthetic organisms that do not grow or grow very slowly under laboratory conditions, which is usually the case for those living in extreme habitats.

The present protocol describes step-by-step autofluorescence imaging and evaluation of phycobiliprotein changes in red algae based on spectral analysis. This is a label-free and non-destructive method to evaluate cellular adaptation to extreme habitats, when only scarce material is available and cells grow slowly, or not at all, under laboratory conditions.

Red algae (Rhodophyta) contain phycobiliproteins and colonize habitats with dim light, however some (e.g., some Chroothece species) can also develop in ...

Quantification of Mitochondrial Membrane Potential and Superoxide Levels

Fluorescence-Based Quantification of Mitochondrial Membrane Potential and Superoxide Levels Using Live Imaging in HeLa Cells

Mitochondria are dynamic organelles critical for metabolic homeostasis by controlling energy production via ATP synthesis. To support cellular metabolism, various mitochondrial quality control mechanisms cooperate to maintain a healthy mitochondrial network. One such pathway is mitophagy, where PTEN-induced kinase 1 (PINK1) and Parkin phospho-ubiquitination of damaged mitochondria facilitate autophagosome sequestration and subsequent removal from the cell via lysosome fusion. Mitophagy is important for cellular homeostasis, and mutations in Parkin are linked to Parkinson's disease (PD). Due to these findings, there has been a significant emphasis on investigating mitochondrial damage and turnover to understand the molecular mechanisms and dynamics of mitochondrial quality control. Here, live-cell imaging was used to visualize the mitochondrial network of HeLa cells, to quantify the mitochondrial membrane potential and superoxide levels following treatment with carbonyl cyanide m-chlorophenyl hydrazone (CCCP), a mitochondrial uncoupling agent. In addition, a PD-linked mutation of Parkin (ParkinT240R) that inhibits Parkin-dependent mitophagy was expressed to determine how mutant expression impacts the mitochondrial network compared to cells expressing wild-type Parkin. The protocol outlined here describes a simple workflow using fluorescence-based approaches to quantify mitochondrial membrane potential and superoxide levels effectively.

Author Spotlight: Fluorescence-Based Quantification of Mitochondrial Membrane Potential and Superoxide Levels Using Live Imaging in HeLa Cells  

This technique describes an effective workflow to visualize and quantitatively measure mitochondrial membrane potential and superoxide levels within HeLa cells using fluorescence-based live imaging.

Mitochondria are dynamic organelles critical for metabolic homeostasis by controlling energy production via ATP synthesis. To support cellular metabolism, ...