The authors thank Sharmin Hossain for the time-lapse movie capturing growth of an immature neuron and Derek Dunfield for the electron microscopy images of SCE micromicropipette tips.

Equipment set-up:
<ol>
<li>The electrical equipment required for this technique are a stimulator, an oscilloscope, and a micropipette holder fitted with a silver wire to be inserted into the micropipette. We typically use an Axon instruments Axoporator 800A stimulator, however other common stimulators such as the Grass instruments SD9 stimulators are also suitable for single-cell electroporation. The stimulator is connected to the headstage, which interfaces with the silver wire electrode to be placed inside the glass micropipette containing an internal conducting solution. The other lead from the stimulator is connected to the oscilloscope input, which also receives input from the tadpole external ground. The tadpole external ground is simply a silver wire that remains in electrotonic contact with the tadpole during electroporation. The circuit is complete once the micropipette is placed in contact with the tadpole. </li>
<li>Single cell electroporation requires a microscope with a good working distance (at least 5cm), since this gives room to maneuver and allows the micropipette to be brought in at an angle of about 30-45 degrees. </li>
</ol>
Fabrication of micropipettes:
Preparation of appropriate micropipettes is a critical step in this protocol. The difficulty lies in balancing a narrow tip (less than 1&mu;m in diameter) with one that will not easily break when puncturing the tissue. We have also found that tips with a taper angle of greater than 10 degrees are preferable.
<ol>
<li>Prepare a pulled glass micropipette with a tip diameter of less than 1&mu;m, and a tip taper angle of greater than 10 degrees. <ol>
<li>Borosilicate glass with an internal filament with outer diameter of 1.5mm and inner diameter of 0.75mm is well suited for this purpose. <ol>
<li>The relatively thick glass provides a degree of rigidity to the tip.</li>
<li>The internal filament is important in drawing back-filled fluid towards the micropipette tip.</li>
</ol></li>
</ol></li>
<li>Fabrication of suitable tips will often require several modifications based on observed results. Different tissues may require a tip with a slightly different shape or tip diameter.</li>
</ol>
Single-cell electroporation protocol:
For new users, it is advisable to begin with fluorescent dextran dyes since, unlike genetically encoded fluorescent proteins, they can be immediately seen under epifluorescence. Use of fluorescent dyes enables the user to directly see the location of the micropipette tip within the tissue, and upon electroporation gives instant feedback as to whether the equipment has been properly set up and whether the micropipette tip is appropriate.
<ol>
<li>Fill micropipette with solution of compound to be electroporated either by using a loading syringe or by back-filling. <ol>
<li>If using plasmid DNA, ensure that samples are endotoxin-free, and at a concentration between 1-2&micro;g/ml, prepared in water.</li>
<li>The concentration of fluorescent dextran dyes should be empirically determined. We often use fluorescent dyes diluted in water to approximately 300&micro;M.</li>
<li>Typically, volumes of 0.5-1&micro;l are sufficient.</li>
<li>Prior to loading, briefly centrifuge solutions at high speed to reduce the amount of particulate debris drawn into the micropipette, which may lead to clogging.</li>
<li>Air bubbles within the tip often affect current flow and should be dislodged by vigorously flicking the micropipette prior to mounting on the headstage.</li>
</ol></li>
<li>Insert micropipette into holder or headstage mounted on a 3-axis micromanipulator. Ensure that silver wire electrode is inserted into micropipette, and is in electrotonic contact with the internal solution.</li>
<li>Anesthetize tadpoles by immersion in 0.02% MS-222 (3-aminobenzoic acid ethyl ester) prepared in tadpole rearing medium (Steinberg's solution, pH 7.4). <ol>
<li>We normally use stage 44-48 albino Xenopus laevis tadpoles in our experiments.</li>
<li>Tadpoles are usually fully anesthetized after approximately 5 minutes.</li>
<li>Several tadpoles may be anesthetized simultaneously to reduce wait-time, however, avoid exposure to MS-222 for periods beyond 1 hour.</li>
</ol></li>
<li>Transfer a single anesthetized tadpole to electroporation chamber using a plastic transfer micropipette. <ol>
<li>The electroporation chamber can be easily made in-house by carving a tadpole-shaped cavity in a small Sylgard&reg; silicone block, and inserting a silver ground wire into the cavity such that it will be in electrotonic contact with the tadpole in the chamber.</li>
</ol></li>
<li>Position tadpole dorsal side up using a moistened paintbrush.</li>
<li>Lower the micropipette until it is in contact with the skin, directly adjacent to the region of interest. <ol>
<li>Targeting regions with densely packed cell bodies will greatly increase chances of successful electroporation. We typically target the tadpole optic tectum.</li>
<li>Bring the micropipette in at an angle of 30 to 45 degrees. Shallower trajectories make skin puncture more difficult, while steeper angles often obstruct one's view.</li>
</ol></li>
<li>Continued lowering of the micropipette will initially dimple the skin, and then pass through into the underlying tissue. <ol>
<li>Superficial cells are preferred for in vivo imaging, and can be targeted by ensuring that the micropipette tip is lowered slowly upon dimpling of skin.</li>
</ol></li>
<li>One should constantly monitor the micropipette resistance once placed within the tissue. If using an Axoporator 800A (Axon Instruments), resistances of approximately 10-40M&Omega; are optimal. Resistance is a good indicator of whether the tip diameter is appropriate for single-cell electroporation. <ol>
<li>High micropipette resistances (&gt;100M&Omega;) are indicative of clogged tips or micropipettes with tips that are too narrow.</li>
<li>Low micropipette resistances (&lt;10M&Omega;) are indicative of a tip diameter that is too large, often a result of a broken tip.</li>
<li>If using a stimulator that does not provide a direct resistance measure, micropipette resistance can indirectly measured by observing the amplitude of the voltage pulses measured by the oscilloscope (discussed below).</li>
</ol></li>
<li>Apply voltage train. <ol>
<li>For plasmid DNA, pulse trains found to be most effective consist of negative square wave pulses 1ms in duration, delivered at 300Hz with pulse train duration of 500ms. <ol>
<li>The pulse voltage is determined empirically such that the amplitude of pulses measured on the oscilloscope is between 0.75 and 1.5&mu;A.</li>
</ol></li>
<li>For fluorescent dextran dyes, positive square wave pulses of 300&mu;s duration are delivered at a frequency of 300Hz with pulse train duration of 10ms. <ol>
<li>As with the protocol for DNA, the pulse voltage is determined empirically such that the amplitude of pulses measured on the oscilloscope is between 0.75 and 1.5&mu;A.</li>
<li>Additional adjustments to the stimulus parameters can be made based on observing the results of electroporation under epifluorescence. <ol>
<li>If electroporated cells appear dim, pulse parameters should be gradually increased until cells appear brighter.</li>
<li>On the other hand, if clusters of cells are labeled, pulse parameters should be reduced until single cells are labeled.</li>
</ol></li>
</ol></li>
</ol></li>
<li>Retract micropipette, re-insert into a different site within the tissue, and apply voltage train. <ol>
<li>To improve yield, we typically electroporate multiple sites in each hemisphere of the tadpole optic tectum with sufficient spacing to ensure that labeled cells do not overlap.</li>
</ol></li>
<li>Transfer the electroporated tadpole into a container with Steinberg's rearing solution. <ol>
<li>Tadpoles will rapidly recover from anesthesia within a few minutes.</li>
</ol></li>
<li>The same micropipette can be used for numerous tadpoles. It is important to constantly observe the amplitude of the pulses observed on the oscilloscope. <ol>
<li>If the pulse amplitude remains low after significantly increasing the pulse intensity this is likely a result of the tip being clogged, often seen when electroporating many tadpoles in one sitting. <ol>
<li>A method to dislodge a minor tip clog is to apply a single positive pulse after inserting the micropipette into the tissue.</li>
<li>If the clog cannot be dislodged, or clogging frequently recurs, replace the micropipette.</li>
</ol></li>
<li>If the pulse amplitude remains very large after significantly decreasing the pulse parameters, this is a sign that the tip has broken and needs to be replaced.</li>
</ol></li>
</ol>
Screening for successfully electroporated cells:
<ol>
<li>After electroporation, tadpoles are screened for labeled cells using an upright epifluorescence microscope.</li>
<li>In the case of fluorescent dyes, tadpoles may be screened after as little as 30 minutes post-electroporation. <ol>
<li>We have found, however, that longer intervals are associated with lower background fluorescence levels.</li>
</ol></li>
<li>For genetically encoded fluorescent proteins, screening is usually conducted at least 12 hours post-electroporation. <ol>
<li>Cells expressing fluorescent proteins will continue to get brighter with time, therefore dim cells may be re-screened after an additional interval of 12-24 hours.</li>
</ol></li>
<li>Tadpoles are anesthetized as described above, placed in a Sylgard&reg; chamber and coverslipped. Individual tadpoles are then examined under epifluorescence. <ol>
<li>Note that excessive exposure to epifluorescence light will result in phototoxicity that may kill the labeled cell. Therefore, minimize the amount of time that cells are exposed to the fluorescent light.</li>
</ol></li>
<li>Return tadpoles to container with Steinberg's rearing solution.</li>
</ol>

<ol>
	<li>Bestman, J. E., Ewald, R. C., Chiu, S. L., Cline, H. T. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=In+vivo+single-cell+electroporation+for+transfer+of+DNA+and+macromolecules.">In vivo single-cell electroporation for transfer of DNA and macromolecules.</a> Nature Protocols. 1, 1267-1272 (2006).</li><li>Dunfield, D., Haas, K. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Single+cell+electroporation.">Single cell electroporation.</a> Encyclopedia of Neuroscience (4th Ed.). , .</li><li>Haas, K., Jensen, K., Sin, W. C., Foa, L., Cline, H. T. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Targeted+electroporation+in+Xenopus+tadpoles+in+vivo--from+single+cells+to+the+entire+brain.">Targeted electroporation in Xenopus tadpoles in vivo--from single cells to the entire brain.</a> Differentiation. 70, 148-154 (2002).</li><li>Haas, K., Sin, W. C., Javaherian, A., Cline, H. T. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Single-cell+electroporation+for+gene+transfer+in+vivo.">Single-cell electroporation for gene transfer in vivo.</a> Neuron. 29, 583-591 (2001).</li></ol>

Single-cell electroporation (SCE) is a powerful tool for determining gene function and performing targeted genetic manipulation. The transparency of the albino tadpole and the accessibility of the brain make this model system ideally suited for visualizing neuronal growth and intracellular events within a live organism. SCE allows the visualization of the growth of a single neuron, and to perform cell-autonomous manipulations within an otherwise unaltered brain. While this video article demonstrates the procedure for sin...

single cell electroporation in vivo within the intact developing brain

Single-cell electroporation (SCE) is a specialized technique allowing the delivery of DNA or other macromolecules into individual cells within intact tissue, including in vivo preparations. The distinct advantage of this technique is that experimental manipulations may be performed on individual cells while leaving the surrounding tissue unaltered, thereby distinguishing cell-autonomous effects from those resulting from global treatments. When combined with advanced in vivo imaging techniques, SCE of fluorescent markers permits direct visualization of cellular morphology, cell growth, and intracellular events over timescales ranging from seconds to days. While this technique is used in a variety of in vivo and ex vivo preparations, we have optimized this technique for use in Xenopus laevis tadpoles. In this video article, we detail the procedure for SCE of a fluorescent dye or plasmid DNA into neurons within the intact brain of the albino Xenopus tadpole. We also discuss methods to optimize yield, and show examples of live two-photon fluorescence imaging of neurons fluorescently labeled by SCE.

Watch this Scientific Journal Video about Single Cell Electroporation in vivo within the Intact Developing Brain at JoVE.com

Single Cell Electroporation in vivo within the Intact Developing Brain

Single-cell electroporation (SCE) is a specialized technique allowing delivery of DNA or other macromolecules into individual cells within intact tissue, including in vivo preparations. Here we detail the procedure for SCE of a fluorescent dye or plasmid DNA into neurons within the intact brain of the Xenopus laevis tadpole.

Single-cell electroporation (SCE) is a specialized technique allowing the delivery of DNA or other macromolecules into individual cells within intact ...

single-cell-electroporation-in-vivo-within-the-intact-developing-brain

Research

JoVE Journal

Biology

13.2K 조회수.  University of British Columbia - UBC. 단일 셀 electroporation (SCE)는 생체내 준비를 포함한 그대로 조직 내의 개별 세포로 DNA 또는 기타 macromolecules의 전달을 허용하는 전문 기술입니다. Xenopus laevis 올챙이의 손상 뇌 내의 뉴런에 형광 염료 또는 플라스미드 DNA의 SCE에 대해 우리가 세부 절차를.

비디오: Single Cell Electroporation in vivo within the Intact Developing Brain

Electroporation of Mycobacteria

High efficiency transformation is a major limitation in the study of mycobacteria. The genus Mycobacterium can be difficult to transform; this is mainly caused by the thick and waxy cell wall, but is compounded by the fact that most molecular techniques have been developed for distantly-related species such as Escherichia coli and Bacillus subtilis. In spite of these obstacles, mycobacterial plasmids have been identified and DNA transformation of many mycobacterial species have now been described. The most successful method for introducing DNA into mycobacteria is electroporation. Many parameters contribute to successful transformation; these include the species/strain, the nature of the transforming DNA, the selectable marker used, the growth medium, and the  conditions for the electroporation pulse. Optimized methods for the transformation of both slow- and fast-grower are detailed here. Transformation efficiencies for different mycobacterial species and with various selectable markers are reported.

Mycobacterial pathogenic strategies remain poorly understood. The slow growth rate of most species, the impenetrable nature of the cell-wall, and the hazards of working with pathogens make mycobacteria difficult to study and are largely responsible for our poor understanding of these organisms. In this video we will demonstrate the technique of electroporation, which involves subjecting cells to a brief high electrical impulse to allow the entry of DNA. It is the most widely used method for introducing DNA into mycobacterial cells.

High efficiency transformation is a major limitation in the study of mycobacteria. The genus Mycobacterium can be difficult to transform; this is mainly ...

In utero and ex vivo Electroporation for Gene Expression in Mouse Retinal Ganglion Cells

The retina and its sole output neuron, the retinal ganglion cell (RGC), comprise an excellent model in which to examine biological questions such as cell differentiation, axon guidance, retinotopic organization and synapse formation[1]. One drawback is the inability to efficiently and reliably manipulate gene expression in RGCs in vivo, especially in the otherwise accessible murine visual pathways. Transgenic mice can be used to manipulate gene expression, but this approach is often expensive, time consuming, and can produce unwanted side effects. In chick, in ovo electroporation is used to manipulate gene expression in RGCs for examining retina and RGC development. Although similar electroporation techniques have been developed in neonatal mouse pups[2], adult rats[3], and embryonic murine retinae in vitro[4], none of these strategies allow full characterization of RGC development and axon projections in vivo. To this end, we have developed two applications of electroporation, one in utero and the other ex vivo, to specifically target embryonic murine RGCs[5, 6]. 
With in utero retinal electroporation, we can misexpress or downregulate specific genes in RGCs and follow their axon projections through the visual pathways in vivo, allowing examination of guidance decisions at intermediate targets, such as the optic chiasm, or at target regions, such as the lateral geniculate nucleus. Perturbing gene expression in a subset of RGCs in an otherwise wild-type background facilitates an understanding of gene function throughout the retinal pathway. Additionally, we have developed a companion technique for analyzing RGC axon growth in vitro. We electroporate embryonic heads ex vivo, collect and incubate the whole retina, then prepare explants from these retinae several days later. Retinal explants can be used in a variety of in vitro assays in order to examine the response of electroporated RGC axons to guidance cues or other factors. In sum, this set of techniques enhances our ability to misexpress or downregulate genes in RGCs and should greatly aid studies examining RGC development and axon projections.

In utero and ex vivo Electroporation for Gene Expression in Mouse Retinal Ganglion Cells

Here we present two techniques for manipulating gene expression in murine retinal ganglion cells (RGCs) by in utero and ex vivo electroporation. These techniques enable one to examine how alterations in gene expression affect RGC development, axon guidance, and functional properties.

The retina and its sole output neuron, the retinal ganglion cell (RGC), comprise an excellent model in which to examine biological questions such as cell ...

DNA Transfection of Mammalian Skeletal Muscles using In Vivo Electroporation

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, &beta;2a-DHPR, etc. ; structural proteins, i.e. minidystrophin and &alpha;-actinin; and membrane proteins, i.e. &alpha;1s-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.

DNA Transfection of Mammalian Skeletal Muscles using In Vivo Electroporation

We describe detailed procedures for the efficient transfection of plasmid DNA into the fibers of foot muscles of live mice using electroporation and the subsequent visualization of protein expression using fluorescence microscopy.

 A growing interest in cell biology is to express transgenically modified forms of essential proteins (e.g. fluorescently tagged constructs and/or mutant ...

Single-cell Suction Recordings from Mouse Cone Photoreceptors

Rod and cone photoreceptors in the retina are responsible for light detection. In darkness, cyclic nucleotide-gated (CNG) channels in the outer segment are open and allow cations to flow steadily inwards across the membrane, depolarizing the cell. Light exposure triggers the closure of the CNG channels, blocks the inward cation current flow, and thus results in cell hyperpolarization. Based on the polarity of photoreceptors, a suction recording method was developed in 1970s that, unlike the classic patch-clamp technique, does not require penetrating the plasma membrane 1. Drawing the outer segment into a tightly-fitting glass pipette filled with extracellular solution allows recording the current changes in individual cells upon test-flash exposure. However, this well-established "outer-segment-in (OS-in)" suction recording is not suitable for mouse cone recordings, because of the low percentage of cones in the mouse retina (3%) and the difficulties in identifying the cone outer segments. Recently, an inner-segment-in (IS-in) recording configuration was developed to draw the inner segment/nuclear region of the photoreceptor into the recording pipette 2,3. In this video, we will show how to record from individual mouse cone photoresponses using single-cell suction electrode.

We will show how to record flash responses from single mouse cones using a suction electrode.

Rod and cone photoreceptors in the retina are responsible for light detection. In darkness, cyclic nucleotide-gated (CNG) channels in the outer segment ...

In vivo Imaging of Intact Drosophila Larvae at Sub-cellular Resolution

Recent improvements in optical imaging, genetically encoded fluorophores and genetic tools allowing efficient establishment of desired transgenic animal lines have enabled biological processes to be studied in the context of a living, and in some instances even behaving, organism. In this protocol we will describe how to anesthetize intact Drosophila larvae, using the volatile anesthetic desflurane, to follow the development and plasticity of synaptic populations at sub-cellular resolution1-3. While other useful methods to anesthetize Drosophila melanogaster larvae have been previously described4,5,6,7,8, the protocol presented herein demonstrates significant improvements due to the following combined key features: (1) A very high degree of anesthetization; even the heart beat is arrested allowing for lateral resolution of up to 150 nm1, (2) a high survival rate of &gt; 90% per anesthetization cycle, permitting the recording of more than five time-points over a period of hours to days2 and (3) a high sensitivity enabling us in 2 instances to study the dynamics of proteins expressed at physiological levels. In detail, we were able to visualize the postsynaptic glutamate receptor subunit GluR-IIA expressed via the endogenous promoter1 in stable transgenic lines and the exon trap line FasII-GFP1. (4) In contrast to other methods4,7 the larvae can be imaged not only alive, but also intact (i.e. non-dissected) allowing observation to occur over a number of days1. The accompanying video details the function of individual parts of the in vivo imaging chamber2,3, the correct mounting of the larvae, the anesthetization procedure, how to re-identify specific positions within a larva and the safe removal of the larvae from the imaging chamber.

In vivo Imaging of Intact Drosophila Larvae at Sub-cellular Resolution

This protocol describes a reliable method for anesthetization and imaging of intact Drosophila melanogaster larvae. We have utilized the volatile anesthetic desflurane to allow for repetitive imaging at sub-cellular resolution and re-identification of structures for up to a few days1.

Recent improvements in optical imaging, genetically encoded fluorophores and genetic tools allowing efficient establishment of desired transgenic animal ...

In vivo Electroporation of Morpholinos into the Adult Zebrafish Retina

Many devastating inherited eye diseases result in progressive and irreversible blindness because humans cannot regenerate dying or diseased retinal neurons. In contrast, the adult zebrafish retina possesses the robust ability to spontaneously regenerate any neuronal class that is lost in a variety of different retinal damage models, including retinal puncture, chemical ablation, concentrated high temperature, and intense light treatment 1-8. Our lab extensively characterized regeneration of photoreceptors following constant intense light treatment and inner retinal neurons after intravitreal ouabain injection 2, 5, 9. In all cases, resident M&uuml;ller glia re-enter the cell cycle to produce neuronal progenitors, which continue to proliferate and migrate to the proper retinal layer, where they differentiate into the deficient neurons. 
	We characterized five different stages during regeneration of the light-damaged retina that were highlighted by specific cellular responses. We identified several differentially expressed genes at each stage of retinal regeneration by mRNA microarray analysis 10. Many of these genes are also critical for ocular development. To test the role of each candidate gene/protein during retinal regeneration, we needed to develop a method to conditionally limit the expression of a candidate protein only at times during regeneration of the adult retina. 
	Morpholino oligos are widely used to study loss of function of specific proteins during the development of zebrafish, Xenopus, chick, mouse, and tumors in human xenografts 11-14. These modified oligos basepair with complementary RNA sequence to either block the splicing or translation of the target RNA. Morpholinos are stable in the cell and can eliminate or "knockdown" protein expression for three to five days 12. 
	Here, we describe a method to efficiently knockdown target protein expression in the adult zebrafish retina. This method employs lissamine-tagged antisense morpholinos that are injected into the vitreous of the adult zebrafish eye. Using electrode forceps, the morpholino is then electroporated into all the cell types of the dorsal and central retina. Lissamine provides the charge on the morpholino for electroporation and can be visualized to assess the presence of the morpholino in the retinal cells. 
	Conditional knockdown in the retina can be used to examine the role of specific proteins at different times during regeneration. Additionally, this approach can be used to study the role of specific proteins in the undamaged retina, in such processes as visual transduction and visual processing in second order neurons.

In vivo Electroporation of Morpholinos into the Adult Zebrafish Retina

A method to conditionally knockdown a target protein&#x2019;s expression in the adult zebrafish retina is described, which involves intravitreally injecting antisense morpholinos and electroporating them into the retina. The resulting protein is knocked down for several days, which allows testing the protein&#x2019;s role in the regenerating or intact retina.

Many devastating inherited eye diseases result in progressive and irreversible blindness because humans cannot regenerate dying or diseased retinal ...

In vivo Electroporation of Morpholinos into the Regenerating Adult Zebrafish Tail Fin

Certain species of urodeles and teleost fish can regenerate their tissues. Zebrafish have become a widely used model to study the spontaneous regeneration of adult tissues, such as the heart1, retina2, spinal cord3, optic nerve4, sensory hair cells5, and fins6.
The zebrafish fin is a relatively simple appendage that is easily manipulated to study multiple stages in epimorphic regeneration. Classically, fin regeneration was characterized by three distinct stages: wound healing, blastema formation, and fin outgrowth. After amputating part of the fin, the surrounding epithelium proliferates and migrates over the wound. At 33 &deg;C, this process occurs within six hours post-amputation (hpa, Figure 1B)6,7. Next, underlying cells from different lineages (ex. bone, blood, glia, fibroblast) re-enter the cell cycle to form a proliferative blastema, while the overlying epidermis continues to proliferate (Figure 1D)8. Outgrowth occurs as cells proximal to the blastema re-differentiate into their respective lineages to form new tissue (Figure 1E)8. Depending on the level of the amputation, full regeneration is completed in a week to a month.
The expression of a large number of gene families, including wnt, hox, fgf, msx, retinoic acid, shh, notch, bmp, and activin-betaA genes, is up-regulated during specific stages of fin regeneration9-16. However, the roles of these genes and their encoded proteins during regeneration have been difficult to assess, unless a specific inhibitor for the protein exists13, a temperature-sensitive mutant exists or a transgenic animal (either overexpressing the wild-type protein or a dominant-negative protein) was generated7,12. We developed a reverse genetic technique to quickly and easily test the function of any gene during fin regeneration.
Morpholino oligonucleotides are widely used to study loss of specific proteins during zebrafish, Xenopus, chick, and mouse development17-19. Morpholinos basepair with a complementary RNA sequence to either block pre-mRNA splicing or mRNA translation. We describe a method to efficiently introduce fluorescein-tagged antisense morpholinos into regenerating zebrafish fins to knockdown expression of the target protein. The morpholino is micro-injected into each blastema of the regenerating zebrafish tail fin and electroporated into the surrounding cells. Fluorescein provides the charge to electroporate the morpholino and to visualize the morpholino in the fin tissue.
This protocol permits conditional protein knockdown to examine the role of specific proteins during regenerative fin outgrowth. In the Discussion, we describe how this approach can be adapted to study the role of specific proteins during wound healing or blastema formation, as well as a potential marker of cell migration during blastema formation.

In vivo Electroporation of Morpholinos into the Regenerating Adult Zebrafish Tail Fin

We describe a method to conditionally knockdown the expression of a target protein during adult zebrafish fin regeneration. This technique involves micro-injecting and electroporating antisense oligonucleotide morpholinos into fin tissue, which allows testing the protein&#x2019;s role in various stages of fin regeneration, including wound healing, blastema formation, and regenerative outgrowth.

Certain species of urodeles and teleost fish can regenerate their tissues. Zebrafish have become a widely used model to study the spontaneous regeneration ...

Confocal Imaging of Single Mitochondrial Superoxide Flashes in Intact Heart or In Vivo

Mitochondrion is a critical intracellular organelle responsible for energy production and intracellular signaling in eukaryotic systems. Mitochondrial dysfunction often accompanies and contributes to human disease. Majority of the approaches that have been developed to evaluate mitochondrial function and dysfunction are based on in vitro or ex vivo measurements. Results from these experiments have limited ability in determining mitochondrial function in vivo. Here, we describe a novel approach that utilizes confocal scanning microscopy for the imaging of intact tissues in live aminals, which allows the evaluation of single mitochondrial function in a real-time manner in vivo. First, we generate transgenic mice expressing the mitochondrial targeted superoxide indicator, circularly permuted yellow fluorescent protein (mt-cpYFP). Anesthetized mt-cpYFP mouse is fixed on a custom-made stage adaptor and time-lapse images are taken from the exposed skeletal muscles of the hindlimb. The mouse is subsequently sacrificed and the heart is set up for Langendorff perfusion with physiological solutions at 37 &deg;C. The perfused heart is positioned in a special chamber on the confocal microscope stage and gentle pressure is applied to immobilize the heart and suppress heart beat induced motion artifact. Superoxide flashes are detected by real-time 2D confocal imaging at a frequency of one frame per second. The perfusion solution can be modified to contain different respiration substrates or other fluorescent indicators. The perfusion can also be adjusted to produce disease models such as ischemia and reperfusion. This technique is a unique approach for determining the function of single mitochondrion in intact tissues and in vivo.

Confocal Imaging of Single Mitochondrial Superoxide Flashes in Intact Heart or In Vivo

Confocal scanning microscopy is applied for imaging single mitochondrial events in perfused heart or skeletal muscles in live animal. Real-time monitoring of single mitochondrial processes such as superoxide flashes and membrane potential fluctuations enables the evaluation of mitochondrial function in a physiologically relevant context and during pathological perturbations.

Mitochondrion is a critical intracellular organelle responsible for energy production and intracellular signaling in eukaryotic systems. Mitochondrial ...

Assessment of Calcium Sparks in Intact Skeletal Muscle Fibers

Maintaining homeostatic Ca2+ signaling is a fundamental physiological process in living cells. Ca2+ sparks are the elementary units of Ca2+ signaling in the striated muscle fibers that appear as highly localized Ca2+ release events mediated by ryanodine receptor (RyR) Ca2+ release channels on the sarcoplasmic reticulum (SR) membrane. Proper assessment of muscle Ca2+ sparks could provide information on the intracellular Ca2+&#160;handling properties of healthy and diseased striated muscles. Although Ca2+ sparks events are commonly seen in resting cardiomyocytes, they are rarely observed in resting skeletal muscle fibers; thus there is a need for methods to generate and analyze sparks in skeletal muscle fibers.
Detailed here is an experimental protocol for measuring Ca2+ sparks in isolated flexor digitorm brevis (FDB) muscle fibers using fluorescent Ca2+ indictors and laser scanning confocal microscopy. In this approach, isolated FDB fibers are exposed to transient hypoosmotic stress followed by a return to isotonic physiological solution. Under these conditions, a robust Ca2+ sparks response is detected adjacent to the sarcolemmal membrane in young healthy FDB muscle fibers. Altered Ca2+ sparks response is detected in dystrophic or aged skeletal muscle fibers. This approach has recently demonstrated that membrane-delimited signaling involving cross-talk between inositol (1,4,5)-triphosphate receptor (IP3R) and RyR contributes to Ca2+ spark activation in skeletal muscle. In summary, our studies using osmotic stress induced Ca2+ sparks showed that this intracellular response reflects a muscle signaling mechanism in physiology and aging/disease states, including mouse models of muscle dystrophy (mdx mice) or amyotrophic lateral sclerosis (ALS model).

Described here is a method to directly measure calcium sparks, the elementary units of Ca2+ release from sarcoplasmic reticulum&#160;in intact skeletal muscle fibers. This method utilizes osmotic-stress-mediated triggering of Ca2+ release from ryanodine receptor in isolated muscle fibers. The dynamics and homeostatic capacity of intracellular Ca2+ signaling can be employed to assess muscle function in health and disease.

Maintaining homeostatic Ca2+ signaling is a fundamental physiological process in living cells. Ca2+ sparks are the elementary units of Ca2+ signaling in ...

In Vivo Microinjection and Electroporation of Mouse Testis

This video and article contribution gives a comprehensive description of microinjection and electroporation of mouse testis in vivo. This particular transfection technique for testicular mouse cells allows the study of unique processes in spermatogenesis.
The following protocol focuses on transfection of testicular mouse cells with plasmid constructs. Specifically, we used the reporter vector pEGFP-C1, which expresses enhanced green fluorescent protein (eGFP) and also the pDsRed2-N1 vector expressing red fluorescent protein (DsRed2). Both encoded reporter genes were under the control of the human cytomegalovirus immediate-early promoter (CMV).
For performing gene transfer into mouse testes, the reporter plasmid constructs are injected into testes of living mice. To that end, the testis of an anaesthetized animal is exposed and the site of microinjection is prepared. Our preferred place of injection is the efferent duct, with the ultimately connected rete testis as the anatomical transport route of the spermatozoa between the testis and the epididymis. In this way, the filling of the seminiferous tubules after microinjection is excellently managed and controlled due to the use of stained DNA solutions. After observing a sufficient filling of the testis by its colored tubule structure, the organ is electroporated. This enables the transfer of the DNA solution into the testicular&#160;cells. Following 3 days of incubation, the testis is removed and investigated under the microscope for green or red&#160;fluorescence, illustrating transfection success.
Generally, this protocol can be employed for delivering DNA- or RNA- constructs into living mouse testis in order to (over)express or knock down genes, facilitating in vivo gene function analysis. Furthermore, it is suitable for studying reporter constructs or putative gene regulatory elements. Thus, the main advantages of the electroporation technique are fast performance in combination with low effort as well as the moderate technical equipment and skills required compared to alternative techniques.

In Vivo Microinjection and Electroporation of Mouse Testis

This article describes microinjection and electroporation of mouse testis in&#160;vivo as a&#160;transfection technique for testicular mouse cells to study unique processes of spermatogenesis. The presented protocol involves steps of glass capillary preparation, microinjection via the efferent duct, and transfection by electroporation.

This video and article contribution gives a comprehensive description of microinjection and electroporation of mouse testis in vivo. This particular ...