We thank Dr. T. Otis, Department of Neurobiology, UCLA, for sharing the TPLSM facility with us, Dr. C. Fahlke, Institute of Physiology, RWTH Aachen, Germany, for the kind donation of the pEYFP-ClC1 plasmid, and Mr. R. Serrano for technical support. This work was supported by grants from NIH/NIAMS grants AR047664 and AR54816.

Experimental Procedures for in vivo electroporation of FDB and IO muscles
<ol>
<li>Before starting the in vivo electroporation protocols, mammalian expression plasmids must be amplified to yield concentration in the range of to 2-5 &mu;g plasmid/&mu;l of TE. Note: We routinely use commercial amplification kits and follow the manufacturer&#x2019; procedures. Commercial expression plasmids carrying the CMV promoter work very well in skeletal muscle in vivo transfections. </li>
<li>Aliquot the necessary volume (10-20 &mu;l) of the plasmid solution and save it in two 0.5 ml Eppendorf tubes (one for each foot of the mouse).</li>
<li>Prepare a solution containing 2 mg/ml hyaluronidase in sterile Tyrode.</li>
<li>Using an anesthetizing box, deeply anesthetize a mouse using 4% isoflurane in O2 with an approved gas anesthetic machine. Place the animal on a heating pad (37 &deg;C) and maintain the anesthesia using a rodent face mask. Monitor the anesthetic depth by toe pinch reflex.</li>
<li>Under observation with a dissection microscope, inject 10 &mu;L of the hyaluronidase solution under the footpads of one foot of the mouse using a 1&#x201D; long 33 gauge sterile needle. Penetrate the skin at a point close to the heel of the foot and advance the needle subcutaneously towards the base of the toes for ~1/4&#x201D;.</li>
<li>Repeat the procedure with the other foot if so desired.</li>
<li>Disconnect the anesthesia and place the mouse in a cage. Allow it to fully recover from anesthesia.</li>
<li>After one hour, anesthetize the animal for a second time and place it on the heating pad. Following the same procedure described for the hyaluronidase solution, inject a total of 20-50 &mu;g of the plasmid DNA (depending on the size of the plasmid construct). The total injection volume should be less than 20 &mu;L/foot. Note: when 15-20 &mu;L is necessary, it is advisable to close the skin at the needle entry point with tissue-glue.</li>
<li>Disconnect the anesthesia and place the mouse in a cage. Allow it to fully recover from anesthesia and wait for 10-15 min.</li>
<li>Anesthetize the animal for the third time and place it on the heating pad.</li>
<li>Select one foot of the animal. Place one gold-plated acupuncture needle under the skin at heel, and a second one at the base of the toes. Electrodes are oriented parallel to each other and perpendicular to the long axis of the foot.</li>
<li>Connect the head of the needles (electrodes) to the electrical stimulator using micro-clip connectors. Electroporate the muscles by applying 20 pulses, 20 ms in duration/each, at 1Hz. Depending on the spacing of the electrodes, the pulses&#x2019; voltage amplitude is adjusted (by monitoring with an oscilloscope) to yield an electric field of ~100 V/cm. Note: No contractions in response to the stimuli should be observed if the level of anesthesia is adequate. </li>
<li>If so desired, repeat the above procedures in the contralateral foot of the animal.</li>
<li>Return the animal to its cage and once fully recovered from anesthesia maintain it under observation. Note: if the procedure went normally, the animal should regain full mobility within 30 minutes and afterwards is ready to be sent back to the animal room at the vivarium. The injections of hyaluronidase and DNA in the footpads do not have noticeable adverse effects on the animals.
Once recovered from anesthesia, mice are able to amble normally around the cage. As an additional precaution, add to the drinking water of the animal Carprofen at 0.0027 mg/ml for 2 days as an analgesic.</li>
<li>Protein expression can be assayed 2-8 days after transfection. However, sustained expression of many proteins has been observed for months.</li>
</ol>
NOTE: All animal procedures were approved by the UCLA Chancellor's Animal Research Committee as mandated by the Animal Welfare Act and the PHS Policy on Humane Care and Use of Laboratory Animals.
Representative Results:
The correct implementation of the in vivo electroporation procedures described above should result in the effective transfection of plasmids in FDB and IO muscles. However, the efficiency of expression of transgenic protein variants will depend on the plasmid, the size and complexity of the protein, the functional properties of the protein, and a number of other variables out of our control. As illustrated in Figure 1 for an FDB muscle electroporated in the presence of a commercial plasmid (pmR-mCherry) encoding for mCherry protein, most of the muscle fibers are transfected with our protocol as illustrated by the fact the most of them display red fluorescence. This does not rule out the possibility that individual fibers exhibit different degrees of protein expression, or that few fibers are not transfected at all 3. It should be noted that the efficient expression of large amounts of fluorescent proteins such as mCherry, EGFP, ECFP, and EYFP, does not impair the muscle fibers&#x2019; excitability and excitation-contraction coupling properties. In fact, they are indistinguishable from those in sham transfected muscles (results not shown).
Practical approaches used to verify the localized expression of fluorescently-tagged proteins in skeletal muscle fibers.
The intracellular localization of the di novo expressed transgenic proteins is routinely evaluated by simultaneous acquiring, using the TPLSM, fluorescent images of the respective tags and images of well-identified markers of cellular structures. The most typical of these latter are: a) second harmonic generation (SHG) images, which arise from the myosin anisotropy of the sarcomeric A bands (centered at the M-lines)9,10 and, b) di-8-ANEPPS fluorescence images, which can be obtained by labeling the surface and transverse tubular (T-tubule) system membranes of the muscle fibers with this impermeant potentiometric dye11. In fluorescence images of muscle fibers stained with di-8-ANEPPS, the T-tubules appear as narrow bands of fluorescence oriented approximately orthogonal to the long axis of the fiber. These bands are unequally spaced from each other: they are separated by a long distance which spans across the M-lines, and a short one that spans across the Z-line11.
Expression and localization of &alpha;-actinin-EGFP in skeletal muscle fibers.
An example of the expression of a tagged variant of the structural muscle protein &alpha;-actinin is shown in Figure 2. This protein is known to be a major component of the Z-line and, as such, is routinely used as a marker of this structure12. We transfected FDB and IO muscles with the plasmid pEGFPN1-&alpha;-actinin1 encoding for human (non-muscle) &alpha;-actinin tagged at the C terminal with EGFP. Six days after transfection, we found that, as assessed by EGFP fluorescence distribution, &alpha;-actinin is mostly expressed at narrow bands equally spaced along the fiber axis. A single band per sarcomere is seen (Figures 2A &amp; 2D). The colocalization of these bands with the Z-lines is demonstrated by comparing the distribution of EGFP fluorescence with the SHG (Figure 2B) and di-8-ANEPPS (Figure 2E) images. As shown by the overlay image (Figure 2C), &alpha;-actinin-EGFP bands alternate with the SHG bands, indicating that they are located midway between two consecutive M-bands, coinciding with the location of Z-lines. In transfected muscle fibers stained with di-8-ANEPPS, &alpha;-actinin-EGFP bands are seen centered between every pair of T-tubules (Figure 2F) which are known to flank the Z-lines (i.e. separated by a shorter distance), thus indicating that transgenic &alpha;-actinin is targeted to the Z-line.
Expression of DHPR&alpha;1s tagged at the N-terminus with EGFP
The efficiency of muscle transfection with pEGFPC1.1-DHPR &alpha;1s is verified in TPLSM images (e.g. Figure 3A) showing that most fibers express EGFP-DHPR&alpha;1s (a transmembrane protein). The most prominent feature of transfected fibers is the double-banded pattern of EGFP fluorescence (Figures 3A &amp; 3B) as would be expected if this protein was targeted to the T-tubules. It can also be observed in Figure 3 that while different fibers display various levels of fluorescence intensity, the banded fluorescence pattern of an individual fiber seems to be maintained homogeneous along the fiber. At higher magnification (Figure 3B), it can be clearly seen that the uneven spacing among bands is similar to that observed in fibers stained di-8-ANEPPS (e.g. Figure 2E). The overlay images (Figure 3B &amp; 3E) illustrate that the SHG bands are located within the larger spacing between EGFP-DHPR&alpha;1s bands, corroborating that this protein is at the T-tubules. Additional FRET measurements with the non-fluorescent lipophilic anion DPA- (data not shown) further demonstrate that the EGFP moiety is within few nanometers of the inner leaflet of the T-tubules&#x2019; membranes.
Localization and functional evaluation of the expression of EYFP-ClC1
Fibers transfected with pEYFP-ClC1, which encodes an EYFP-tagged construct (at the N-terminal) of the skeletal muscle chloride channel (ClC1), display EYFP fluorescence bands (Figure 4A) with a pattern similar to that observed for EGFP-DHPR&alpha;1s&#x2019; expression (Figure 3A) and corresponding to the T-tubule arrangement as illustrated with di-8-ANEPPS staining (Figure 2E). As expected, the superimposition of EYFP-ClC1 (Figure 4A) and SHG images (Figure 4B) as shown in the overlay (Figure 4C), illustrate that the SHG bands are centered at the large spacing of between EYFP fluorescence bands.
In order to assess whether the expression of EYFP-ClC1 results in a significant increase in the resting conductance of the muscle fibers, as expected from the overexpression of functional chloride channels, we enzymatically dissociated muscle fibers from the transfected FDB muscle and studied their electrophysiological properties using a two-microelectrodes experimental setup described previously6,11,13. Figure 5 shows results from two fibers: one expressing large amounts of EYFP-ClC1 as assessed from global fluorescence intensity measurements in a standard fluorescence microscope (not shown), and the other is a non-transfected control. The voltage record in Figure 5A (obtained from the muscle fiber expressing EYFP-ClC1) demonstrates that due to the excessive resting conductance, the fiber is almost non-excitable. Current stimulus pulses of up to 400nA (0.5ms) needed to be used in order to elicit a very small active response (no overshoot, Figure 5A). After replacement of the external Tyrode solution to one containing 500 &mu;M of the chloride channel blocker 9-ACA, 200 nA current pulses were sufficient to elicit an action potential (Figure 5B), although much slower and broader than those recorded in the control fiber (Figure 5C). As expected, addition of 9-ACA had minimal effects on this control fiber (Figure 5D).
<img src="/files/ftp_upload/1520/1520fig1tmb.jpg" alt="Figure 1" border="0" /> Figure 1. Transfection efficiency of the in-vivo electroporation method. Brightfield (A) and fluorescence (B) images of an FDB muscle transfected with pmR-mCherry. Days after electroporation protocol: 12 days. Please <a href="https://www.jove.com/files/ftp_upload/1520/1520fig1.jpg">click here</a> for a larger version of figure 1.
<img src="/files/ftp_upload/1520/1520figure2.jpg" alt="Figure 2" /> Figure 2. Expression and targeting of &alpha;-actinin-EGFP in FDB fibers. Panels A and B are EGFP fluorescence and SHG images, respectively, of a fiber expressing &alpha;-actinin-EGFP. Panel C is a superposition of the images in A and B. Panels D and E are fluorescence images of another fiber expressing &alpha;-actinin-EGFP and stained with di-8-ANEPPS, respectively. Panel F is the superposition of images in D and E. Days after electroporation protocol: 6 days.
<img src="/files/ftp_upload/1520/1520figure3.jpg" alt="Figure 3" /> Figure 3. Expression and targeting of EGFP-DHPR&alpha;1s in FDB fibers. Panel A, EGFP fluorescence image of a group of fibers expressing EGFP-DHPR&alpha;1s. Panel B is an enlargement of the square in panel A in order to better show the banded pattern of the protein expression. Panel C is the SHG image corresponding to the image in panel A. Panel D is the overlay of the images A and C. Panel E is an enlargement of the area indicated in panel D. Days after electroporation protocol: 20 days. 
<img src="/files/ftp_upload/1520/1520figure4.jpg" alt="Figure 4" /> Figure 4. Expression and targeting of EYFP-ClC1 in FDB fibers. Panels A and B are EYFP fluorescence and SHG images, respectively, of fibers expressing EYFP-ClC1. Panel C is the superposition of the images in panels A and B. Panel D is an intensity profile measured along the white line highlighted in the image in panel C. Days after electroporation protocol: 7 days.
<img src="/files/ftp_upload/1520/1520figure5.jpg" alt="Figure 5" /> Figure 5. Electrophysiological assessment fibers expressing transgenic EYFP-ClC1. Panels A and B are voltage records from a fiber expressing EYFP-ClC1 in response to current pulses before and after treatment with 9-ACA, respectively. Panels C and D are voltage records (action potentials) elicited in response to current pulses in a non-transfected fiber before and after treatment with 9-ACA, respectively.

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The authors have nothing to disclose.

We describe here the detailed steps that should be followed in order to attain effective transfections of DNA plasmids into skeletal muscle fibers by in vivo electroporation. The main advantages of our approach are the simplicity of implementation, and its minimal invasiveness which results in negligible health hazard to the animals. In reality, the above described electroporation protocols do not entail much more than two subcutaneous injections per foot, followed by an electrical stimulation protocol, all of w...

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.

Watch this Scientific Journal Video about DNA Transfection of Mammalian Skeletal Muscles using In Vivo Electroporation at JoVE.com

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.

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 ...

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17.7K Views.  David Geffen School of Medicine, University of California, Los Angeles. 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. 

Video: DNA Transfection of Mammalian Skeletal Muscles using In Vivo Electroporation

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.

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 ...

Intranuclear Microinjection of DNA into Dissociated Adult Mammalian Neurons

Primary neuronal cell cultures are valuable tools to study protein function since they represent a more biologically relevant system compared to immortalized cell lines. However, the post-mitotic nature of primary neurons prevents effective heterologous protein expression using common procedures such as electroporation or chemically-mediated transfection. Thus, other techniques must be employed in order to effectively express proteins in these non-dividing cells.
In this article, we describe the steps required to perform intranuclear injections of cDNA constructs into dissociated adult sympathetic neurons. This technique, which has been applied to different types of neurons, can successfully induce heterologous protein expression. The equipment essential for the microinjection procedure includes an inverted microscope to visualize cells, a glass injection pipet filled with cDNA solution that is connected to a N2(g) pressure delivery system, and a micromanipulator. The micromanipulator coordinates the injection motion of microinjection pipet with a brief pulse of pressurized N2 to eject cDNA solution from the pipet tip. 
This technique does not have the toxicity associated with many other transfection methods and enables multiple DNA constructs to be expressed at a consistent ratio. The low number of injected cells makes the microinjection procedure well suited for single cell studies such as electrophysiological recordings and optical imaging, but may not be ideal for biochemical assays that require a larger number of cells and higher transfection efficiencies. Although intranuclear microinjections require an investment of equipment and time, the ability to achieve high levels of heterologous protein expression in a physiologically relevant environment makes this technique a very useful tool to investigate protein function.

Direct intranuclear injection of cDNA is an effective transfection technique for post-mitotic cells. This method provides high levels of heterologous protein expression from single or multiple cDNA constructs and enables protein function to be studied in a physiologically relevant environment with a variety of single cell assays.

Primary neuronal cell cultures are valuable tools to study protein function since they represent a more biologically relevant system compared to ...

Analysis of DNA Double-strand Break (DSB) Repair in Mammalian Cells

DNA double-strand breaks are the most dangerous DNA lesions that may lead to massive loss of genetic information and cell death. Cells repair DSBs using two major pathways: nonhomologous end joining (NHEJ) and homologous recombination (HR). Perturbations of NHEJ and HR are often associated with premature aging and tumorigenesis, hence it is important to have a quantitative way of measuring each DSB repair pathway. Our laboratory has developed fluorescent reporter constructs that allow sensitive and quantitative measurement of NHEJ and HR. The constructs are based on an engineered GFP gene containing recognition sites for a rare-cutting I-SceI endonuclease for induction of DSBs. The starting constructs are GFP negative as the GFP gene is inactivated by an additional exon, or by mutations. Successful repair of the I-SceI-induced breaks by NHEJ or HR restores the functional GFP gene. The number of GFP positive cells counted by flow cytometry provides quantitative measure of NHEJ or HR efficiency.

Analysis of DNA Double-strand Break DSB Repair in Mammalian Cells

This article describes GFP-based fluorescence in vivo assays that separately quantify homologous recombination and nonhomologous end joining in mammalian cells.

DNA double-strand breaks are the most dangerous DNA lesions that may lead to massive loss of genetic information and cell death.  Cells repair DSBs using ...

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 ...

Ex Vivo Assessment of Contractility, Fatigability and Alternans in Isolated Skeletal Muscles

Described here is a method to measure contractility of isolated skeletal muscles. Parameters such as muscle force, muscle power, contractile kinetics, fatigability, and recovery after fatigue can be obtained to assess specific aspects of the excitation-contraction coupling (ECC) process such as excitability, contractile machinery and Ca2+ handling ability. This method removes the nerve and blood supply and focuses on the isolated skeletal muscle itself. We routinely use this method to identify genetic components that alter the contractile property of skeletal muscle though modulating Ca2+ signaling pathways. Here, we describe a newly identified skeletal muscle phenotype, i.e., mechanic alternans, as an example of the various and rich information that can be obtained using the in vitro muscle contractility assay. Combination of this assay with single cell assays, genetic approaches and biochemistry assays can provide important insights into the mechanisms of ECC in skeletal muscle.

Ex Vivo Assessment of Contractility, Fatigability and Alternans in Isolated Skeletal Muscles

We describe a method to directly measure muscle force, muscle power, contractile kinetics and fatigability of isolated skeletal muscles in an in vitro system using field stimulation. Valuable information on Ca2+ handling properties and contractile machinery of the muscle can be obtained using different stimulating protocols.

 
Described here is a method to measure contractility of isolated skeletal muscles. Parameters such as muscle force, muscle power, contractile kinetics, ...

Real-time Imaging of Single Engineered RNA Transcripts in Living Cells Using Ratiometric Bimolecular Beacons

The growing realization that both the temporal and spatial regulation of gene expression can have important consequences on cell function has led to the development of diverse techniques to visualize individual RNA transcripts in single living cells. One promising technique that has recently been described utilizes an oligonucleotide-based optical probe, ratiometric bimolecular beacon (RBMB), to detect RNA transcripts that were engineered to contain at least four tandem repeats of the RBMB target sequence in the 3&#8217;-untranslated region. RBMBs are specifically designed to emit a bright fluorescent signal upon hybridization to complementary RNA, but otherwise remain quenched. The use of a synthetic probe in this approach allows photostable, red-shifted, and highly emissive organic dyes to be used for imaging. Binding of multiple RBMBs to the engineered RNA transcripts results in discrete fluorescence spots when viewed under a wide-field fluorescent microscope. Consequently, the movement of individual RNA transcripts can be readily visualized in real-time by taking a time series of fluorescent images. Here we describe the preparation and purification of RBMBs, delivery into cells by microporation and live-cell imaging of single RNA transcripts.

Ratiometric bimolecular beacons (RBMBs) can be used to image single engineered RNA transcripts in living cells. Here, we describe the preparation and purification of RBMBs, delivery of RBMBs into cells by microporation and fluorescent imaging of single RNA transcripts in real-time.

The growing realization that both the temporal and spatial regulation of gene expression can have important consequences on cell function has led to the ...

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 ...

Detection and Analysis of DNA Damage in Mouse Skeletal Muscle In Situ Using the TUNEL Method

Terminal deoxynucleotidyl transferase (TdT) deoxyuridine triphosphate (dUTP) nick end labeling (TUNEL) is the method of using the TdT enzyme to covalently attach a tagged form of dUTP to 3&#8217; ends of double- and single-stranded DNA breaks in cells. It is a reliable and useful method to detect DNA damage and cell death in situ. This video describes dissection, tissue processing, sectioning, and fluorescence-based TUNEL labeling of mouse skeletal muscle. It also describes a method of semi-automated TUNEL signal quantitation. Inherent normal tissue features and tissue processing conditions affect the ability of the TdT enzyme to efficiently label DNA. Tissue processing may also add undesirable autofluorescence that will interfere with TUNEL signal detection. Therefore, it is important to empirically determine tissue processing and TUNEL labeling methods that will yield the optimal signal-to-noise ratio for subsequent quantitation. The fluorescence-based assay described here provides a way to exclude autofluorescent signal by digital channel subtraction. The TUNEL assay, used with appropriate tissue processing techniques and controls, is a relatively fast, reproducible, quantitative method for detecting apoptosis in tissue. It can be used to confirm DNA damage and apoptosis as pathological mechanisms, to identify affected cell types, and to assess the efficacy of therapeutic treatments in vivo.

Detection and Analysis of DNA Damage in Mouse Skeletal Muscle In Situ Using the TUNEL Method

This video describes dissection, tissue processing, sectioning, and fluorescence-based TUNEL labeling of mouse skeletal muscle. It also describes a method for semi-automated analysis of TUNEL labeling.

Terminal deoxynucleotidyl transferase (TdT) deoxyuridine triphosphate (dUTP) nick end labeling (TUNEL) is the method of using the TdT enzyme to covalently ...

DNA Electroporation, Isolation and Imaging of Myofibers

Mature muscle has a unique structure that is amenable to live cell imaging. Herein, we describe the experimental protocol for expressing fluorescently labeled proteins in the flexor digitorum brevis (FDB) muscle. Conditions have been optimized to provide a large number of high quality myofibers expressing the electroporated plasmid while minimizing muscle damage. The method employs fluorescent tags on various proteins. Combining this expression method with high resolution confocal microscopy permits live cell imaging, including imaging after laser-induced damage. Fluorescent dyes combined with imaging of fluorescently-tagged proteins provides information regarding the basic structure of muscle and its response to stimuli.

This protocol utilizes electroporation to introduce and express fluorescently labeled proteins in mouse muscle fibers. Following recovery after electroporation, fibers are isolated. Individual fibers are then imaged using high resolution confocal microscopy to visualize muscle structure.

Mature muscle has a unique structure that is amenable to live cell imaging. Herein, we describe the experimental protocol for expressing fluorescently ...

Examination of Proteins Bound to Nascent DNA in Mammalian Cells Using  BrdU-ChIP-Slot-Western Technique

Histone deacetylases 1 and 2 (HDAC1,2) localize to the sites of DNA replication. In the previous study, using a selective inhibitor and a genetic knockdown system, we showed novel functions for HDAC1,2 in replication fork progression and nascent chromatin maintenance in mammalian cells. Additionally, we used a BrdU-ChIP-Slot-Western technique that combines chromatin immunoprecipitation (ChIP) of bromo-deoxyuridine (BrdU)-labeled DNA with slot blot and Western analyses to quantitatively measure proteins or histone modification associated with nascent DNA.
Actively dividing cells were treated with HDAC1,2 selective inhibitor or transfected with siRNAs against Hdac1 and Hdac2 and then newly synthesized DNA was labeled with the thymidine analog bromodeoxyuridine (BrdU). The BrdU labeling was done at a time point when there was no significant cell cycle arrest or apoptosis due to the loss of HDAC1,2 functions. Following labeling of cells with BrdU, chromatin immunoprecipitation (ChIP) of histone acetylation marks or the chromatin-remodeler was performed with specific antibodies. BrdU-labeled input DNA and the immunoprecipitated (or ChIPed) DNA was then spotted onto a membrane using the slot blot technique and immobilized using UV. The amount of nascent DNA in each slot was then quantitatively assessed using Western analysis with an anti-BrdU antibody. The effect of loss of HDAC1,2 functions on the levels of newly synthesized DNA-associated histone acetylation marks and chromatin remodeler was then determined by normalizing the BrdU-ChIP signal obtained from the treated samples to the control samples.

In this protocol, we describe a novel BrdU-ChIP-Slot-Western technique to examine proteins and histone modifications associated with newly synthesized or nascent DNA.