Cardiomyocytes, the workhorse cell of the heart, contain exquisitely organized cytoskeletal and contractile elements that generate the contractile force used to pump blood. Individual cardiomyocytes were first isolated over 40 years ago in order to better study the physiology and structure of heart muscle. Techniques have rapidly improved to include enzymatic digestion via coronary perfusion. More recently, analyzing the contractility and calcium flux of isolated myocytes has provided a vital tool in the cellular and sub-cellular analysis of heart failure. Echocardiography and EKGs provide information about the heart at an organ level only. Cardiomyocyte cell culture systems exist, but cells lack physiologically essential structures such as organized sarcomeres and t-tubules required for myocyte function within the heart. In the protocol presented here, cardiomyocytes are isolated via Langendorff perfusion. The heart is removed from the mouse, mounted via the aorta to a cannula, perfused with digestion enzymes, and cells are introduced to increasing calcium concentrations. Edge and sarcomere detection software is used to analyze contractility, and a calcium binding fluorescent dye is used to visualize calcium transients of electrically paced cardiomyocytes; increasing understanding of the role cellular changes play in heart dysfunction. Traditionally used to test drug effects on cardiomyocytes, we employ this system to compare myocytes from WT mice and mice with a mutation that causes dilated cardiomyopathy. This protocol is unique in its comparison of live cells from mice with known heart function and known genetics. Many experimental conditions are reliably compared, including genetic or environmental manipulation, infection, drug treatment, and more. Beyond physiologic data, isolated cardiomyocytes are easily fixed and stained for cytoskeletal elements. Isolating cardiomyocytes via perfusion is an extremely versatile method, useful in studying cellular changes that accompany or lead to heart failure in a variety of experimental conditions.
18 Related JoVE Articles!
Analysis of Cardiomyocyte Development using Immunofluorescence in Embryonic Mouse Heart
Institutions: Northwestern University, University of California, San Francisco.
During heart development, the generation of myocardial-specific structural and functional units including sarcomeres, contractile myofibrils, intercalated discs, and costameres requires the coordinated assembly of multiple components in time and space. Disruption in assembly of these components leads to developmental heart defects. Immunofluorescent staining techniques are used commonly in cultured cardiomyocytes to probe myofibril maturation, but this ex vivo
approach is limited by the extent to which myocytes will fully differentiate in culture, lack of normal in vivo
mechanical inputs, and absence of endocardial cues. Application of immunofluorescence techniques to the study of developing mouse heart is desirable but more technically challenging, and methods often lack sufficient sensitivity and resolution to visualize sarcomeres in the early stages of heart development. Here, we describe a robust and reproducible method to co-immunostain multiple proteins or to co-visualize a fluorescent protein with immunofluorescent staining in the embryonic mouse heart and use this method to analyze developing myofibrils, intercalated discs, and costameres. This method can be further applied to assess cardiomyocyte structural changes caused by mutations that lead to developmental heart defects.
Developmental Biology, Issue 97, Immunofluorescence, mouse, embryonic heart, cardiomyocyte, development, sarcomere, intercalated disc, costamere, s-α-actinin, cryosection
Isolation and Culture of Neonatal Mouse Cardiomyocytes
Institutions: King’s College London, University of California San Diego .
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.
Cellular Biology, Issue 79, Biomedical Engineering, Bioengineering, Molecular Biology, Cell Culture Techniques, Primary Cell Culture, Cell Culture Techniques, Primary Cell Culture, Cell Culture Techniques, Primary Cell Culture, Cell Culture Techniques, Disease Models, Animal, Models, Cardiovascular, Cell Biology, neonatal mouse, cardiomyocytes, isolation, culture, primary cells, NMC, heart cells, animal model
Live Cell Imaging of Primary Rat Neonatal Cardiomyocytes Following Adenoviral and Lentiviral Transduction Using Confocal Spinning Disk Microscopy
Institutions: Max-Planck-Institute for Molecular Biomedicine and Institute of Cell Biology, Yale Cardiovascular Research Center and Section of Cardiovascular Medicine.
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’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.
Cellular Biology, Issue 88, live cell imaging, cardiomyocyte, primary cell culture, adenovirus, lentivirus, confocal spinning disk microscopy
Analysis of Tubular Membrane Networks in Cardiac Myocytes from Atria and Ventricles
Institutions: Heart Research Center Goettingen, University Medical Center Goettingen, German Center for Cardiovascular Research (DZHK) partner site Goettingen, University of Maryland School of Medicine.
In cardiac myocytes a complex network of membrane tubules - the transverse-axial tubule system (TATS) - controls deep intracellular signaling functions. While the outer surface membrane and associated TATS membrane components appear to be continuous, there are substantial differences in lipid and protein content. In ventricular myocytes (VMs), certain TATS components are highly abundant contributing to rectilinear tubule networks and regular branching 3D architectures. It is thought that peripheral TATS components propagate action potentials from the cell surface to thousands of remote intracellular sarcoendoplasmic reticulum (SER) membrane contact domains, thereby activating intracellular Ca2+
release units (CRUs). In contrast to VMs, the organization and functional role of TATS membranes in atrial myocytes (AMs) is significantly different and much less understood. Taken together, quantitative structural characterization of TATS membrane networks in healthy and diseased myocytes is an essential prerequisite towards better understanding of functional plasticity and pathophysiological reorganization. Here, we present a strategic combination of protocols for direct quantitative analysis of TATS membrane networks in living VMs and AMs. For this, we accompany primary cell isolations of mouse VMs and/or AMs with critical quality control steps and direct membrane staining protocols for fluorescence imaging of TATS membranes. Using an optimized workflow for confocal or superresolution TATS image processing, binarized and skeletonized data are generated for quantitative analysis of the TATS network and its components. Unlike previously published indirect regional aggregate image analysis strategies, our protocols enable direct characterization of specific components and derive complex physiological properties of TATS membrane networks in living myocytes with high throughput and open access software tools. In summary, the combined protocol strategy can be readily applied for quantitative TATS network studies during physiological myocyte adaptation or disease changes, comparison of different cardiac or skeletal muscle cell types, phenotyping of transgenic models, and pharmacological or therapeutic interventions.
Bioengineering, Issue 92, cardiac myocyte, atria, ventricle, heart, primary cell isolation, fluorescence microscopy, membrane tubule, transverse-axial tubule system, image analysis, image processing, T-tubule, collagenase
High Efficiency Differentiation of Human Pluripotent Stem Cells to Cardiomyocytes and Characterization by Flow Cytometry
Institutions: Medical College of Wisconsin, Stanford University School of Medicine, Medical College of Wisconsin, Hong Kong University, Johns Hopkins University School of Medicine, Medical College of Wisconsin.
There is an urgent need to develop approaches for repairing the damaged heart, discovering new therapeutic drugs that do not have toxic effects on the heart, and improving strategies to accurately model heart disease. The potential of exploiting human induced pluripotent stem cell (hiPSC) technology to generate cardiac muscle “in a dish” for these applications continues to generate high enthusiasm. In recent years, the ability to efficiently generate cardiomyogenic cells from human pluripotent stem cells (hPSCs) has greatly improved, offering us new opportunities to model very early stages of human cardiac development not otherwise accessible. In contrast to many previous methods, the cardiomyocyte differentiation protocol described here does not require cell aggregation or the addition of Activin A or BMP4 and robustly generates cultures of cells that are highly positive for cardiac troponin I and T (TNNI3, TNNT2), iroquois-class homeodomain protein IRX-4 (IRX4), myosin regulatory light chain 2, ventricular/cardiac muscle isoform (MLC2v) and myosin regulatory light chain 2, atrial isoform (MLC2a) by day 10 across all human embryonic stem cell (hESC) and hiPSC lines tested to date. Cells can be passaged and maintained for more than 90 days in culture. The strategy is technically simple to implement and cost-effective. Characterization of cardiomyocytes derived from pluripotent cells often includes the analysis of reference markers, both at the mRNA and protein level. For protein analysis, flow cytometry is a powerful analytical tool for assessing quality of cells in culture and determining subpopulation homogeneity. However, technical variation in sample preparation can significantly affect quality of flow cytometry data. Thus, standardization of staining protocols should facilitate comparisons among various differentiation strategies. Accordingly, optimized staining protocols for the analysis of IRX4, MLC2v, MLC2a, TNNI3, and TNNT2 by flow cytometry are described.
Cellular Biology, Issue 91, human induced pluripotent stem cell, flow cytometry, directed differentiation, cardiomyocyte, IRX4, TNNI3, TNNT2, MCL2v, MLC2a
Methods to Assess Subcellular Compartments of Muscle in C. elegans
Institutions: University of Nottingham.
Muscle is a dynamic tissue that responds to changes in nutrition, exercise, and disease state. The loss of muscle mass and function with disease and age are significant public health burdens. We currently understand little about the genetic regulation of muscle health with disease or age. The nematode C. elegans
is an established model for understanding the genomic regulation of biological processes of interest. This worm’s body wall muscles display a large degree of homology with the muscles of higher metazoan species. Since C. elegans
is a transparent organism, the localization of GFP to mitochondria and sarcomeres allows visualization of these structures in vivo
. Similarly, feeding animals cationic dyes, which accumulate based on the existence of a mitochondrial membrane potential, allows the assessment of mitochondrial function in vivo
. These methods, as well as assessment of muscle protein homeostasis, are combined with assessment of whole animal muscle function, in the form of movement assays, to allow correlation of sub-cellular defects with functional measures of muscle performance. Thus, C. elegans
provides a powerful platform with which to assess the impact of mutations, gene knockdown, and/or chemical compounds upon muscle structure and function. Lastly, as GFP, cationic dyes, and movement assays are assessed non-invasively, prospective studies of muscle structure and function can be conducted across the whole life course and this at present cannot be easily investigated in vivo
in any other organism.
Developmental Biology, Issue 93, Physiology, C. elegans, muscle, mitochondria, sarcomeres, ageing
Derivation of Highly Purified Cardiomyocytes from Human Induced Pluripotent Stem Cells Using Small Molecule-modulated Differentiation and Subsequent Glucose Starvation
Institutions: Stanford University School of Medicine, Stanford University School of Medicine.
Human induced pluripotent stem cell-derived cardiomyocytes (hiPSC-CMs) have become an important cell source to address the lack of primary cardiomyocytes available for basic research and translational applications. To differentiate hiPSCs into cardiomyocytes, various protocols including embryoid body (EB)-based differentiation and growth factor induction have been developed. However, these protocols are inefficient and highly variable in their ability to generate purified cardiomyocytes. Recently, a small molecule-based protocol utilizing modulation of Wnt/β-Catenin signaling was shown to promote cardiac differentiation with high efficiency. With this protocol, greater than 50%-60% of differentiated cells were cardiac troponin-positive cardiomyocytes were consistently observed. To further increase cardiomyocyte purity, the differentiated cells were subjected to glucose starvation to specifically eliminate non-cardiomyocytes based on the metabolic differences between cardiomyocytes and non-cardiomyocytes. Using this selection strategy, we consistently obtained a greater than 30% increase in the ratio of cardiomyocytes to non-cardiomyocytes in a population of differentiated cells. These highly purified cardiomyocytes should enhance the reliability of results from human iPSC-based in vitro
disease modeling studies and drug screening assays.
Developmental Biology, Issue 97, Human induced pluripotent stem cells, cardiac differentiation, small molecule compounds, cardiomyocytes, glucose starvation
Measurement of Maximum Isometric Force Generated by Permeabilized Skeletal Muscle Fibers
Institutions: University of Michigan Medical School, University of Michigan Medical School, University of Michigan Medical School, University of Michigan Medical School.
Analysis of the contractile properties of chemically skinned, or permeabilized, skeletal muscle fibers offers a powerful means by which to assess muscle function at the level of the single muscle cell. Single muscle fiber studies are useful in both basic science and clinical studies. For basic studies, single muscle fiber contractility measurements allow investigation of fundamental mechanisms of force production, and analysis of muscle function in the context of genetic manipulations. Clinically, single muscle fiber studies provide useful insight into the impact of injury and disease on muscle function, and may be used to guide the understanding of muscular pathologies. In this video article we outline the steps required to prepare and isolate an individual skeletal muscle fiber segment, attach it to force-measuring apparatus, activate it to produce maximum isometric force, and estimate its cross-sectional area for the purpose of normalizing the force produced.
Bioengineering, Issue 100, Muscle physiology, skeletal muscle, single muscle fiber, permeabilized, cross-sectional area, isometric force, specific force
Preparation of Segmented Microtubules to Study Motions Driven by the Disassembling Microtubule Ends
Institutions: Russian Academy of Sciences, Federal Research Center of Pediatric Hematology, Oncology and Immunology, Moscow, Russia, University of Pennsylvania.
Microtubule depolymerization can provide force to transport different protein complexes and protein-coated beads in vitro
. The underlying mechanisms are thought to play a vital role in the microtubule-dependent chromosome motions during cell division, but the relevant proteins and their exact roles are ill-defined. Thus, there is a growing need to develop assays with which to study such motility in vitro
using purified components and defined biochemical milieu. Microtubules, however, are inherently unstable polymers; their switching between growth and shortening is stochastic and difficult to control. The protocols we describe here take advantage of the segmented microtubules that are made with the photoablatable stabilizing caps. Depolymerization of such segmented microtubules can be triggered with high temporal and spatial resolution, thereby assisting studies of motility at the disassembling microtubule ends. This technique can be used to carry out a quantitative analysis of the number of molecules in the fluorescently-labeled protein complexes, which move processively with dynamic microtubule ends. To optimize a signal-to-noise ratio in this and other quantitative fluorescent assays, coverslips should be treated to reduce nonspecific absorption of soluble fluorescently-labeled proteins. Detailed protocols are provided to take into account the unevenness of fluorescent illumination, and determine the intensity of a single fluorophore using equidistant Gaussian fit. Finally, we describe the use of segmented microtubules to study microtubule-dependent motions of the protein-coated microbeads, providing insights into the ability of different motor and nonmotor proteins to couple microtubule depolymerization to processive cargo motion.
Basic Protocol, Issue 85, microscopy flow chamber, single-molecule fluorescence, laser trap, microtubule-binding protein, microtubule-dependent motor, microtubule tip-tracking
Monitoring Actin Disassembly with Time-lapse Microscopy
Institutions: Harvard Medical School.
Cellular Biology, Issue 1, cytoskeleton, actin, timelapse, filament, chamber
Isolation, Culture, and Functional Characterization of Adult Mouse Cardiomyoctyes
Institutions: Beth Israel Deaconess Medical Center, Harvard Medical School, Sapienza University.
The use of primary cardiomyocytes (CMs) in culture has provided a powerful complement to murine models of heart disease in advancing our understanding of heart disease. In particular, the ability to study ion homeostasis, ion channel function, cellular excitability and excitation-contraction coupling and their alterations in diseased conditions and by disease-causing mutations have led to significant insights into cardiac diseases. Furthermore, the lack of an adequate immortalized cell line to mimic adult CMs, and the limitations of neonatal CMs (which lack many of the structural and functional biomechanics characteristic of adult CMs) in culture have hampered our understanding of the complex interplay between signaling pathways, ion channels and contractile properties in the adult heart strengthening the importance of studying adult isolated cardiomyocytes. Here, we present methods for the isolation, culture, manipulation of gene expression by adenoviral-expressed proteins, and subsequent functional analysis of cardiomyocytes from the adult mouse. The use of these techniques will help to develop mechanistic insight into signaling pathways that regulate cellular excitability, Ca2+
dynamics and contractility and provide a much more physiologically relevant characterization of cardiovascular disease.
Cellular Biology, Issue 79, Medicine, Cardiology, Cellular Biology, Anatomy, Physiology, Mice, Ion Channels, Primary Cell Culture, Cardiac Electrophysiology, adult mouse cardiomyocytes, cell isolation, IonOptix, Cell Culture, adenoviral transfection, patch clamp, fluorescent nanosensor
Capillary Force Lithography for Cardiac Tissue Engineering
Institutions: University of Washington, University of Washington.
Cardiovascular disease remains the leading cause of death worldwide1
. Cardiac tissue engineering holds much promise to deliver groundbreaking medical discoveries with the aims of developing functional tissues for cardiac regeneration as well as in vitro
screening assays. However, the ability to create high-fidelity models of heart tissue has proven difficult. The heart’s extracellular matrix (ECM) is a complex structure consisting of both biochemical and biomechanical signals ranging from the micro- to the nanometer scale2
. Local mechanical loading conditions and cell-ECM interactions have recently been recognized as vital components in cardiac tissue engineering3-5
A large portion of the cardiac ECM is composed of aligned collagen fibers with nano-scale diameters that significantly influences tissue architecture and electromechanical coupling2
. Unfortunately, few methods have been able to mimic the organization of ECM fibers down to the nanometer scale. Recent advancements in nanofabrication techniques, however, have enabled the design and fabrication of scalable scaffolds that mimic the in vivo
structural and substrate stiffness cues of the ECM in the heart6-9
Here we present the development of two reproducible, cost-effective, and scalable nanopatterning processes for the functional alignment of cardiac cells using the biocompatible polymer poly(lactide-co-glycolide) (PLGA)8
and a polyurethane (PU) based polymer. These anisotropically nanofabricated substrata (ANFS) mimic the underlying ECM of well-organized, aligned tissues and can be used to investigate the role of nanotopography on cell morphology and function10-14
Using a nanopatterned (NP) silicon master as a template, a polyurethane acrylate (PUA) mold is fabricated. This PUA mold is then used to pattern the PU or PLGA hydrogel via UV-assisted or solvent-mediated capillary force lithography (CFL), respectively15,16
. Briefly, PU or PLGA pre-polymer is drop dispensed onto a glass coverslip and the PUA mold is placed on top. For UV-assisted CFL, the PU is then exposed to UV radiation (λ = 250-400 nm) for curing. For solvent-mediated CFL, the PLGA is embossed using heat (120 °C) and pressure (100 kPa). After curing, the PUA mold is peeled off, leaving behind an ANFS for cell culture. Primary cells, such as neonatal rat ventricular myocytes, as well as human pluripotent stem cell-derived cardiomyocytes, can be maintained on the ANFS2
Bioengineering, Issue 88, Nanotopography, Anisotropic, Nanofabrication, Cell Culture, Cardiac Tissue Engineering
Laser-inflicted Injury of Zebrafish Embryonic Skeletal Muscle
Institutions: Max Delbrück Center for Molecular Medicine.
Various experimental approaches have been used in mouse to induce muscle injury with the aim to study muscle regeneration, including myotoxin injections (bupivacaine, cardiotoxin or notexin), muscle transplantations (denervation-devascularization induced regeneration), intensive exercise, but also murine muscular dystrophy models such as the mdx
mouse (for a review of these approaches see 1
). In zebrafish, genetic approaches include mutants that exhibit muscular dystrophy phenotypes (such as runzel2
) and antisense oligonucleotide morpholinos that block the expression of dystrophy-associated genes4
. Besides, chemical approaches are also possible, e.g.
with Galanthamine, a chemical compound inhibiting acetylcholinesterase, thereby resulting in hypercontraction, which eventually leads to muscular dystrophy5
. However, genetic and pharmacological approaches generally affect all muscles within an individual, whereas the extent of physically inflicted injuries are more easily controlled spatially and temporally1
. Localized physical injury allows the assessment of contralateral muscle as an internal control. Indeed, we recently used laser-mediated cell ablation to study skeletal muscle regeneration in the zebrafish embryo6
, while another group recently reported the use of a two-photon laser (822 nm) to damage very locally the plasma membrane of individual embryonic zebrafish muscle cells7
Here, we report a method for using the micropoint laser (Andor Technology) for skeletal muscle cell injury in the zebrafish embryo. The micropoint laser is a high energy laser which is suitable for targeted cell ablation at a wavelength of 435 nm. The laser is connected to a microscope (in our setup, an optical microscope from Zeiss) in such a way that the microscope can be used at the same time for focusing the laser light onto the sample and for visualizing the effects of the wounding (brightfield or fluorescence). The parameters for controlling laser pulses include wavelength, intensity, and number of pulses.
Due to its transparency and external embryonic development, the zebrafish embryo is highly amenable for both laser-induced injury and for studying the subsequent recovery. Between 1 and 2 days post-fertilization, somitic skeletal muscle cells progressively undergo maturation from anterior to posterior due to the progression of somitogenesis from the trunk to the tail8, 9
. At these stages, embryos spontaneously twitch and initiate swimming. The zebrafish has recently been recognized as an important vertebrate model organism for the study of tissue regeneration, as many types of tissues (cardiac, neuronal, vascular etc.) can be regenerated after injury in the adult zebrafish10, 11
Developmental Biology, Issue 71, Anatomy, Physiology, Medicine, Molecular Biology, Cellular Biology, Biomedical Engineering, Genetics, Zebrafish, skeletal muscle, cell ablation, injury, regeneration, damage, laser pulses, tissue, embryos, Danio rerio, animal model
Immunostaining of Dissected Zebrafish Embryonic Heart
Institutions: Mayo Clinic College of Medicine.
Zebrafish embryo becomes a popular in vivo
vertebrate model for studying cardiac development and human heart diseases due to its advantageous embryology and genetics 1,2
. About 100-200 embryos are readily available every week from a single pair of adult fish. The transparent embryos that develop ex utero
make them ideal for assessing cardiac defects 3
. The expression of any gene can be manipulated via morpholino technology or RNA injection 4
. Moreover, forward genetic screens have already generated a list of mutants that affect different perspectives of cardiogenesis 5
Whole mount immunostaining is an important technique in this animal model to reveal the expression pattern of the targeted protein to a particular tissue 6
. However, high resolution images that can reveal cellular or subcellular structures have been difficult, mainly due to the physical location of the heart and the poor penetration of the antibodies.
Here, we present a method to address these bottlenecks by dissecting heart first and then conducting the staining process on the surface of a microscope slide. To prevent the loss of small heart samples and to facilitate solution handling, we restricted the heart samples within a circle on the surface of the microscope slides drawn by an immEdge pen. After the staining, the fluorescence signals can be directly observed by a compound microscope.
Our new method significantly improves the penetration for antibodies, since a heart from an embryonic fish only consists of few cell layers. High quality images from intact hearts can be obtained within a much reduced procession time for zebrafish embryos aged from day 2 to day 6. Our method can be potentially extended to stain other organs dissected from either zebrafish or other small animals.
Developmental Biology, Issue 59, Zebrafish, Danio rerio, Embryonic Heart, Cardiology, Dissection, Immunostaining
Visualisation and Quantification of Intracellular Interactions of Neisseria meningitidis and Human α-actinin by Confocal Imaging
Institutions: University of Bristol, UK.
The Opc protein of Neisseria meningitidis
(Nm, meningococcus) is a surface-expressed integral outer membrane protein, which can act as an adhesin and an effective invasin for human epithelial and endothelial cells. We have identified endothelial surface-located integrins as major receptors for Opc, a process which requires Opc to first bind to integrin ligands such as vitronectin and via these to the cell-expressed receptors1
. This process leads to bacterial invasion of endothelial cells2
. More recently, we observed an interaction of Opc with a 100kDa protein found in whole cell lysates of human cells3
. We initially observed this interaction when host cell proteins separated by electrophoresis and blotted on to nitrocellulose were overlaid with Opc-expressing Nm. The interaction was direct and did not involve intermediate molecules. By mass spectrometry, we established the identity of the protein as α-actinin. As no surface expressed α-actinin was found on any of the eight cell lines examined, and as Opc interactions with endothelial cells in the presence of serum lead to bacterial entry into the target cells, we examined the possibility of the two proteins interacting intracellularly. For this, cultured human brain microvascular endothelial cells (HBMECs) were infected with Opc-expressing Nm for extended periods and the locations of internalised bacteria and α-actinin were examined by confocal microscopy. We observed time-dependent increase in colocalisation of Nm with the cytoskeletal protein, which was considerable after an eight hour period of bacterial internalisation. In addition, the use of quantitative imaging software enabled us to obtain a relative measure of the colocalisation of Nm with α-actinin and other cytoskeletal proteins. Here we present a protocol for visualisation and quantification of the colocalisation of the bacterium with intracellular proteins after bacterial entry into human endothelial cells, although the procedure is also applicable to human epithelial cells.
Immunology, Issue 44, Neisseria meningitidis, Opc, α-actinin, colocalization
DNA Transfection of Mammalian Skeletal Muscles using In Vivo Electroporation
Institutions: David Geffen School of Medicine, University of California, Los Angeles.
A growing interest in cell biology is to express transgenically modified forms of essential proteins (e.g. fluorescently tagged constructs and/or mutant variants) in order to investigate their endogenous distribution and functional relevance. An interesting approach that has been implemented to fulfill this objective in fully differentiated cells is the in vivo
transfection of plasmids by various methods into specific tissues such as liver1
, skeletal muscle2,3
, and even the brain4
. We present here a detailed description of the steps that must be followed in order to efficiently transfect genetic material into fibers of the flexor digitorum brevis
(FDB) and interosseus
(IO) muscles of adult mice using an in vivo
electroporation approach. The experimental parameters have been optimized so as to maximize the number of muscle fibers transfected while minimizing tissue damages that may impair the quality and quantity of the proteins expressed in individual fibers. We have verified that the implementation of the methodology described in this paper results in a high yield of soluble proteins, i.e. EGFP and ECFP3
, calpain, FKBP12, β2a-DHPR, etc. ; structural proteins, i.e. minidystrophin and α-actinin; and membrane proteins, i.e. α1s-DHPR, RyR1, cardiac Na/Ca2+
exchanger , NaV1.4 Na channel, SERCA1, etc., when applied to FDB, IO and other muscles of mice and rats. The efficient expression of some of these proteins has been verified with biochemical3
and functional evidence5
. However, by far the most common confirmatory approach used by us are standard fluorescent microscopy and 2-photon laser scanning microscopy (TPLSM), which permit to identify not only the overall expression, but also the detailed intracellular localization, of fluorescently tagged protein constructs. The method could be equally used to transfect plasmids encoding for the expression of proteins of physiological relevance (as shown here), or for interference RNA (siRNA) aiming to suppress the expression of normally expressed proteins (not tested by us yet). It should be noted that the transfection of FDB and IO muscle fibers is particularly relevant for the investigation of mammalian muscle physiology since fibers enzymatically dissociated from these muscles are currently one of the most suitable models to investigate basic mechanisms of excitability and excitation-contraction coupling under current or voltage clamp conditions2,6-8
Cellular Biology, Issue 32, electroporation, skeletal muscle, plasmids, protein expression, mouse, two-photon microscopy, fluorescence, transgenic
Fluorescent Labeling of Drosophila Heart Structures
Institutions: San Diego State University, The Sanford Burnham Institute for Medical Research.
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 α-actinin in larval hearts. Additionally, we perform labeling of F-actin and α-actinin in myosin-GFP expressing adult flies and of α-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.
Cellular Biology, Issue 32, Cardiac, cardiomyopathy, dorsal vessel, fluorescence, staining, GFP, larva, immunohistochemistry, microscopy, imaging
Isolation and Cryopreservation of Neonatal Rat Cardiomyocytes
Institutions: College of Veterinary Medicine, North Carolina State University, University of North Carolina at Chapel Hill, North Carolina State University, Soochow University.
Cell culture has become increasingly important in cardiac research, but due to the limited proliferation of cardiomyocytes, culturing cardiomyocytes is difficult and time consuming. The most commonly used cells are neonatal rat cardiomyocytes (NRCMs), which require isolation every time cells are needed. The birth of the rats can be unpredictable. Cryopreservation is proposed to allow for cells to be stored until needed, yet freezing/thawing methods for primary cardiomyocytes are challenging due to the sensitivity of the cells. Using the proper cryoprotectant, dimethyl sulfoxide (DMSO), cryopreservation was achieved. By slowly extracting the DMSO while thawing the cells, cultures were obtained with viable NRCMs. NRCM phenotype was verified using immunocytochemistry staining for α-sarcomeric actinin. In addition, cells also showed spontaneous contraction after several days in culture. Cell viability after thawing was acceptable at 40-60%. In spite of this, the methods outlined allow one to easily cryopreserve and thaw NRCMs. This gives researchers a greater amount of flexibility in planning experiments as well as reducing the use of animals.
Developmental Biology, Issue 98, Cryopreservation, Cardiomyocytes, Cell-based Assay, Cell Culture, Cardiac Research, In Vitro Assay