The authors would like to thank Karen Lawrence for her help with the section describing construction of the electroporation chamber.

1. Construction of the electroporation chamber
<ol>
 <li>Order a microslide, BTX Model 453 with a 3.2mm gap (Harvard Apparatus #45-0105) (Fig. 2A). The metal rails should be completely sealed to the bottom of the slide.</li>
 <li>Use a Dremel tool to cut a handle off a plastic microcentrifuge tube rack. Cut the handle into 5 small rectangular pieces each with the following dimensions: length 0.8cm, height 0.6cm, width 0.3cm (Fig. 2B). These plastic pieces are reusable spacers that will be used to mold individual wells in the microslide chamber.</li>
 <li>Insert the plastic spacers between the metal rails of the microslide at even intervals. The spacers should fit snugly (Fig. 2C).</li>
 <li>Cut the tip off a P200 pipette tip, fit the tip to a 3ml syringe, and fill the syringe with 100% silicone rubber aquarium sealant. Fill the gaps between the plastic spacers with sealant. Be sure to fill the gaps from the bottom up so that no bubbles form between the sealant plug and the base of the microslide.</li>
 <li>Place a metal rod atop the plastic spacers and fasten it with binder clips, so that the spacers are held in place while the sealant dries (Fig. 2D-E). Let the sealant dry overnight.</li>
 <li>Remove the binder clips, metal rod, and plastic spacers (Fig. 2F). Use a scalpel blade to clean the sealant off the top of the metal rails. Use a dissecting microscope to examine the microslide and ensure that no bubbles are present at the bottom of the silicone dams. Remove any sealant film that may have formed in the wells such that bare metal is exposed inside the wells. Fill one well with water and make sure that the water doesn&#x2019;t leak into the adjacent well(s). Repeat for all wells.</li>
 <li>The finished microslide fits in the plastic dish with the metal poles adjacent to the window in the side of the dish (Fig. 2G); the electrodes will eventually be attached to the metal poles.</li>
</ol>
2. DNA preparation
<ol>
 <li>Add plasmid DNA to a 1.5mL microcentrifuge tube on ice and bring the volume up to 150&mu;l with distilled water. Multiple plasmid species may be combined for co-electroporation (e.g., an experimental DsRed construct and a control GFP construct). When calculating the amount of DNA to add to the tube, keep in mind that the final volume of the DNA aliquot will be 60&mu;l. Typically, each construct is used at a final concentration of 0.5&mu;g/&mu;l.</li>
 <li>Precipitate the DNA by adding 15&mu;l 3M sodium acetate (pH 5.2) and 450&mu;l 100% ethanol. Invert or tap the tube several times to mix.</li>
 <li>Spin down the DNA at 4&deg;C, 13,200 rpm, for 30 minutes. Wash the pellet with 70% ethanol, then spin it down again at 4&deg;C, 13,200 rpm, for 15 minutes. Air-dry the pellet until semi-translucent, about 7 minutes, then resuspend in 54&mu;l sterile water. Add 6&mu;l sterile 10X PBS (pH 7.4) and mix.</li>
</ol>
3. Eye collection
<ol>
 <li>Sterilize the electroporation chamber and all instruments with 70% ethanol. While eye collection and dissection need not be performed in a tissue culture hood, disinfect your gloves and benchtop with ethanol and attempt to maintain sterile conditions throughout the procedure.</li>
 <li>Prepare Petri dishes with dissection medium (1:1 ratio of DMEM:F12, 100U/ml penicillin, 100&mu;g/ml streptomycin, 0.29mg/ml L-glutamine, and 5&mu;g/ml insulin): two 35mm dishes each with 3ml medium and one 60mm dish with 6ml medium. This step should be performed in a tissue culture hood.</li>
 <li>Disinfect the head and neck of a newborn (postnatal day 0) mouse pup with 70% ethanol. Quickly decapitate with scissors and transfer the head to a sterile 100mm dish.</li>
 <li>Cut away the scalp with small scissors to expose the eyes. Use curved forceps to gently scoop the eye out of the orbit, and place the eye in a 35mm dish containing dissection medium. It may be helpful to remove the eyes under a dissecting microscope at low power.</li>
 <li>Repeat steps 3.3 and 3.4 until all eyes have been collected. Keep eyes in dissection medium at room temperature while dissecting. You will need 3-4 eyes per DNA aliquot.</li>
</ol>
4. Retinal dissection
<ol>
 <li>Use 70% ethanol to disinfect both a razor blade and the wrapper of a sterile plastic transfer pipette. Cut the tip off the pipette with the blade so that it can suck up a whole eye. Store the pipette in the plastic wrapper when not in use.</li>
 <li>Transfer one eye from the 35mm dish to the 60mm dish. Under the dissecting microscope at high power, use fine forceps to remove any tissue, such as extraocular muscles and fat, from the surface of the eye. Then, remove the optic nerve by pinching it off at the base.</li>
 <li>In order to isolate the retina, poke a small hole in the sclera at the limbus. Insert one prong from both pairs of forceps into the hole (tangential to the retinal surface) and gently tear open the sclera/RPE. In albino mice, the sclera and RPE appear shiny relative to the retinal tissue, which is a homogeneous matte gray color. In pigmented mice, the RPE is black. Leave the lens in place.</li>
 <li>Use the transfer pipette to move the dissected retina into the other 35mm dish with medium.</li>
 <li>Repeat steps 4.2 to 4.4 until all eyes have been dissected.</li>
 <li>Store the retinas in a 37&deg;C tissue culture incubator until you are ready to electroporate.</li>
</ol>
5. Preparation for electroporation
<ol>
 <li>Prepare 35mm dishes of medium. For each DNA aliquot to be electroporated, you will need one dish of dissection medium and one dish of culture medium (dissection medium plus 10% FBS). Label the dishes appropriately.</li>
 <li>Use a P200 pipette and sterile 1X PBS to wash out the chambers in the electroporation dish. Each chamber holds a volume of 60-100&mu;l. Wash out each chamber three times.</li>
 <li>Fill the chambers with the DNA aliquots. Any unused chambers should be filled with 60&mu;l 1X PBS. Connect the electrodes to the electroporation dish.</li>
 <li>Use the following settings on the electroporator: mode, LV; voltage, 30V; pulse length, 50 msec; number of pulses, 5; interval, 950 msec; polarity, unipolar.</li>
</ol>
6. Electroporation
<ol>
 <li>Use fine forceps to grasp the retinas by the lens and transfer them into the electroporation chambers. Each chamber holds up to 3-4 mouse retinas (Fig. 3).</li>
 <li>Use forceps to line up the retinas such that the lens leans against the metal bar attached to the positive electrode. Clean the forceps with a Kimwipe after each transfer to avoid carry-over of DNA from one chamber to the next.</li>
 <li>Once all retinas are aligned, press &#x201C;Start&#x201D; on the electroporator. Tiny bubbles should form on the metal bar attached to the negative electrode.</li>
 <li>Disconnect the electrodes and turn off the electroporator.</li>
 <li>Use forceps to gently move the retinas away from the chamber walls.</li>
 <li>Use a sterile transfer pipette to transfer the retinas from the chambers into the 35mm dishes containing dissection medium.</li>
 <li>Wash out each chamber three times with sterile 1X PBS, then rinse with sterile water. Spray dish with 70% ethanol.</li>
</ol>
7. Placing retinas on filters for culture
<ol>
 <li>Use a transfer pipette to transfer the retinas into the 35mm dishes containing culture medium.</li>
 <li>Label the wells of a sterile 6-well culture plate and fill each well with 3ml culture medium.</li>
 <li>Use sterile forceps to place round Whatman Nuclepore filters, shiny side up, atop the medium in each well.</li>
 <li>Under a dissecting microscope, use a sterile transfer pipette to transfer the retinas onto the filter, lens-side-down. If the retina lands lens-side-up, pick it up with the pipette and attempt to place it again. Do not place more than 4 retinas on one filter, and make sure that the droplets of medium surrounding each retina remain separate from the other droplets. Note that we culture the retina with the lens intact, although other protocols call for the removal of the lens prior to this step9.</li>
 <li>Place the culture plate in a 37&deg;C tissue culture incubator (5% CO2) and grow for the desired amount of time, typically eight days. In our experience, changing the medium is unnecessary during the eight day culture period, although a medium change may be required for longer culture periods.</li>
</ol>
8. Harvesting and flatmounting fluorescent retinal explants
<ol>
 <li>Replace the culture medium in each well with 4% paraformaldehyde / 1X PBS. If the retinas remain stuck to the filters, use forceps to flip the filters over and gently peel the retinas off the filter. Incubate in paraformaldehyde for 30 min. at room temperature. Protect the retinas from light to avoid bleaching the fluorescence.</li>
 <li>Rinse the retinas twice for 10 minutes in 1X PBS.</li>
 <li>Use a disposable pipette to transfer the retinas onto a glass slide in a small drop of PBS. Under a fluorescent dissecting microscope, use forceps to flip the retinas so that they are electroporated-side-up (i.e., lens-side-down).</li>
 <li>Place glass &#x201C;feet&#x201D; made from crushed coverslips (glass shards about 1-2 mm in diameter) at the corners of the slide; these feet prevent flattening of the retina. Place an intact glass coverslip over the slide so that it covers the retinas and rests on the feet. If necessary, use a pipette to add more PBS between the slide and the coverslip.</li>
</ol>
9. Imaging and quantification of fluorescence in flatmount
<ol>
 <li>Use a fluorescent compound microscope equipped with a monochrome camera to image the flatmounted retina at low power (4X objective) in the red and green channels. All retinas must be imaged with the same exposure time for a given fluorescent channel to enable comparison of fluorescence intensities. Make sure that the pixels are not saturated in any image, or else accurate quantification will be impossible. Export images in grayscale TIFF format.</li>
 <li>Open the image set for one retina (i.e.., red channel and green channel) in ImageJ software (<a href="http://rsbweb.nih.gov/ij/">http://rsbweb.nih.gov/ij/</a>). For the sake of this tutorial, the green fluorescent channel (GFP protein) is the control construct that is constant across all retinas in the experiment. The red fluorescent channel (DsRed protein) is the experimental construct that varies for each set of retinas. The images should be in grayscale.</li>
 <li> In ImageJ, select the control green image and specify a circle of interest with diameter 100 units (Analyze/Tools/ROI manager/More/Specify). Duplicate this circle (ROI manager/Add) to create eight circles total. Move circles 1-5 to select five regions that are uniformly electroporated, avoiding the outer edges of the retina and the region overlying the lens (Fig. 4A). Also, select three regions (circles 6-8) outside the retina/lens to measure background fluorescence. Select the red image, un-check the &#x201C;Show all&#x201D; box in ROI manager, and re-check the box. All eight circles should appear on the red image. De-select all circle coordinates in ROI manager.</li>
 <li>With the red image selected, record the mean pixel value for all circles of interest (ROI manager/Measure); measurements 1-8 should appear, where 1-5 are the red retinal measurements and 6-8 are the red background measurements. Select the green image and record the mean pixel value; measurements 9-16 should appear, where 9-13 are the green retinal measurements and 14-16 are the green background measurements. Copy the measurement data into Excel for analysis (Fig. 5).</li>
 <li>Average the three background measurements in both the red and the green channels. Subtract the red background average from each of the five retinal measurements in the red channel; repeat for the green channel. For each retinal region-of-interest, divide the background-subtracted red measurement by the background-subtracted green measurement in order to normalize the experimental red level to the control green level (Fig. 5).</li>
 <li>Determine the average and standard deviation of all normalized measurements for a given DsRed construct (e.g., 5 measurements per retina times 3 separate retinas). In order to quantitatively compare the results of electroporations carried out on different days, always include a &#x201C;standard&#x201D; DsRed/GFP precipitation in each electroporation set. Relative expression values across experiments can be compared by normalizing to the expression level of this &#x201C;standard.&#x201D;</li>
</ol>
10. Representative Results:
A good electroporation results in expression of the DNA construct(s) across 1/4 to 1/3 of the retinal surface (Fig. 4A). Since rod photoreceptors in particular are efficiently transduced, this technique is ideal for quantifying photoreceptor-specific promoter activity (Fig. 4B). We previously used this approach to quantify a range of promoter variants from the rod-specific Rho and Gnat1 loci10. We found that it is possible to quantify promoter activity over a nearly 300-fold range. 
Figure 5 is a sample dataset from a single electroporated retina. In this particular example, the experimental construct pNrl(1.1kb)-DsRed was measured in the red channel and the control construct pNrl(3.2kb)-GFP was measured in the green channel. A complete dataset for the pNrl(1.1kb)-DsRed construct would consist of 6-9 retinas measured in this manner, and standard deviation would be calculated based on all &#x201C;DsRed normalized to GFP&#x201D; values. If we were comparing the expression level of pNrl(1.1kb)-DsRed to, for example, pNrl(0.8kb)-DsRed, then both constructs would need to be electroporated with the same GFP control (e.g., pNrl(3.2kb)-GFP) and imaged at the same exposure times. It is possible to pool data collected on different days if a standard DsRed/GFP electroporation is performed on each day (e.g., pNrl(3.2kb)-dsRed + pNrl(3.2kb)-GFP). For each experimental construct, the normalized DsRed level would subsequently be normalized to the normalized DsRed level of the &#x201C;standard&#x201D; (pNrl(3.2kb)-dsRed). 
The technique we describe here is primarily useful for quantifying the activity of photoreceptor CREs10,13. Cell-type specific cis-regulatory activity can also be quantified in rarer retinal cell types such as bipolar cells14, but this usually requires that the areas of interest to be quantified be selected in vertical cross-sections rather than in flatmount preparations. The same is true of CREs which drive expression in multiple cell types such as photoreceptors and bipolar cells. The experimental procedures are otherwise similar.
<img src="/files/ftp_upload/2821/2821fig1.jpg" alt="Figure 1"/> 
 Figure 1. Overview of the retinal explant electroporation procedure. First, whole eyes are isolated from postnatal day 0 mouse pups and the retinas are dissected (P1). Second, the retinas are placed in chambers filled with DNA and electroporated (P2). Third, the retinas are placed on filters and cultured for eight days (P3). Fourth, the retinal explants are fixed, mounted on slides, and imaged. Fluorescence intensity is measured with ImageJ software (P4). Fifth, the ImageJ data are processed in a spreadsheet program to quantify the difference in activity of various promoters (P5).
<img src="/files/ftp_upload/2821/2821fig2.jpg" alt="Figure 2"/> 
 Figure 2. Construction of the electroporation dish. A) Unmodified microslide chamber from Harvard Apparatus, BTX model 453 (catalog #45-0105). B) A Dremel tool is used to cut the handle off a plastic tube rack. The handle is cut into rectangular spacers with the following dimensions: length 0.8cm, height 0.6cm, width 0.3cm. C) The plastic spacers are fitted into the microslide chamber at equal intervals. Aquarium sealant is injected into the gaps between the spacers (not shown). D) A metal bar is placed over the spacers. E) The bar and spacers are clamped onto the slide with binder clips to hold everything in place as the sealant dries overnight. F) The spacers are removed and the wells are tested to ensure that they are watertight. G) The finished slide fits into the plastic dish with the metal bars adjacent to the window in the side of the dish.
<img src="/files/ftp_upload/2821/2821fig3.jpg" alt="Figure 3"/> 
 Figure 3. Diagram of the electroporation dish with retinas. The chambers are filled with DNA solutions (up to five different solutions at a time). Retinas are placed in the chambers and oriented so that the lens is leaning against the metal bar connected to the positive electrode; three or four retinas will fit in each of the five chambers. The electrical current will cause the negatively-charged DNA molecules to move into the retinal cells.
<img src="/files/ftp_upload/2821/2821fig4.jpg" alt="Figure 4"/> 
 Figure 4. A) ImageJ measurement of retinal fluorescence levels in flatmount. Grayscale flatmount images in the DsRed (experimental) and GFP (control) channels are opened in ImageJ software; note that these images have been colored for illustrative purposes only. Five measurement circles (1 through 5) are placed over uniformly electroporated regions, avoiding the edges and lens (dotted lines). Three measurement circles (6 through 8) are placed outside the retina to determine background fluorescence levels. B) Cross-sectional images of an electroporated retinal explant at high power. The explant was fixed at postnatal day 8, cryoprotected in 30% sucrose/1X PBS overnight at 4&deg;C, embedded in OCT, and cryo-sectioned at 12&mu;m. The fluorescent constructs pNrl(1.1kb)-DsRed and pNrl(3.2kb)-GFP are expressed in photoreceptor cells in the outer nuclear layer (ONL). INL, inner nuclear layer; GCL, ganglion cell layer.
<img src="/files/ftp_upload/2821/2821fig5.jpg" alt="Figure 5"/> 
 Figure 5. Processing of fluorescence data using Excel. In step 1, the mean pixel value for each measurement circle is copied into the spreadsheet (cells B3-B12, F3-F5, H3-H5). Measurements #1-5 are the DsRed retinal values and #6-8 are the DsRed background values; measurements #9-13 are the GFP retinal values and #14-16 are the GFP background values. Note that measurement #1 and #9 correspond to the same measurement circle, as do measurements #2 and #10, and so on. In step 2, the average background value for the DsRed and GFP channels is calculated (cells F6, H6). In step 3, the average background is subtracted from every retinal measurement (cells C3-C12). In step 4, each background-subtracted DsRed measurement is normalized to its corresponding GFP measurement (cells D3-D7).

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J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=ChIP-Seq+identification+of+weakly+conserved+heart+enhancers.">ChIP-Seq identification of weakly conserved heart enhancers.</a> Nat Genet. 42, 806-810 (2010).</li><li>Visel, A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=ChIP-seq+accurately+predicts+tissue-specific+activity+of+enhancers.">ChIP-seq accurately predicts tissue-specific activity of enhancers.</a> Nature. 457, 854-858 (2009).</li><li>Matsuda, T., Cepko, C. 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No conflicts of interest declared.

Explant electroporation is a simple means of quantifying cis-regulatory activity in the developing mouse retina. Compared to cis-regulatory analysis via mouse transgenesis, electroporation is much cheaper, requiring only newborn mouse pups, DNA, dissecting instruments, and electroporation/tissue culture equipment. It is also far less time consuming: one experiment requires only a few hours of preparation time, a culture period of about eight days, and a few hours at the end of the experiment for imaging and data analysis. Explant electroporation is also superior to cell culture-based cis-regulatory analysis since actual retinal tissue is utilized. The retina develops quite normally in explant culture&#x2014;it forms three distinct cellular layers (outer nuclear layer, inner nuclear layer, and ganglion cell layer), although photoreceptors fail to elaborate outer segments.
A further advantage is that explant electroporation is highly reproducible. The same constructs electroporated into different retinas, even on different days, typically results in the same relative expression levels. Furthermore, since the electroporated plasmids are thought to be maintained episomally in the nucleus and are not incorporated into chromosomes, they do not appear to be subject to the same integration site effects that bedevil cis-regulatory analysis performed in transgenic mice.
Explant electroporation does have several limitations. First, only cells that are still in the cell cycle can be efficiently transduced by electroporation15. At P0, rods and other later-born retinal cell types (bipolar cells, amacrine cells, M ller glia) are the main cell populations targeted by this method. Electroporation of cone photoreceptors by P0 electroporation has been reported16 but the efficiency appears to be low. A second limitation is that explant culture beyond two weeks results in progressive malformation of the retina and is therefore not recommended. If promoter quantification is required at late timepoints, however, an in vivo electroporation9 may be performed, followed by retinal dissection at the desired timepoint, flat-mounting of the dissected retina, and quantification as described in section 9. A third limitation is that this assay is only moderately high-throughput. Unlike cell culture-based assays that can test hundreds of constructs in a single experiment, the technique described in this protocol requires a minimum of one whole mouse retina per construct. Thus, only a couple dozen constructs can reasonably be electroporated in a day.
One additional caveat with respect to the quantification of promoter activity using the present approach is that there is a potential for 'bleed-through' of DsRed fluorescence into the GFP channel, particularly when assaying very strong promoters. The reason for this is that the emission spectrum of DsRed partially overlaps that of GFP. To circumvent this issue, optimized emission filters should be used that minimize the spectral overlap between DsRed and GFP. When such optimized filter sets are not available, another potential solution would be to use a blue-shifted fluorescent protein (e.g., BFP or CFP) in lieu of GFP.

<table border="1">
	<tbody>
 
 <tr>
 	<th>Name of the reagent</th>
 <th>Company</th>
 <th>Catalogue number</th>
 <th>Comments (optional)</th>
 </tr>
 <tr>
 	<td>Electroporation dish, Microslide 453</td>
 <td>BTX Harvard Apparatus</td>
 <td>45-0105</td>
 <td>See protocol section 1 for modifications</td>
 </tr>

 <tr>
 	<td>100% silicone rubber aquarium cement</td>
 <td>Perfecto Manufacturing</td>
 <td>&nbsp;</td>
 <td>&nbsp;</td>
 </tr>

 <tr>
 	<td>Plastic microtube rack</td>
 <td>Fisher Scientific</td>
 <td>05-541</td>
 <td>Only the rack handle will be used</td>
 </tr>

 <tr>
 	<td>Dremel tool</td>
 <td>&nbsp;</td>
 <td>&nbsp;</td>
 <td>For cutting the handle off the plastic tube rack</td>
 </tr>

 <tr>
 	<td>DMEM</td>
 <td>Gibco/Invitrogen</td>
 <td>11965</td>
 <td>&nbsp;</td>
 </tr>

 <tr>
 	<td>F12</td>
 <td>Gibco/Invitrogen</td>
 <td>11765</td>
 <td>&nbsp;</td>
 </tr>

 <tr>
 	<td>L-Glu/pen/strep</td>
 <td>Gibco/Invitrogen</td>
 <td>10378-016</td>
 <td>100X concentration</td>
 </tr>

 <tr>
 	<td>Insulin</td>
 <td>Sigma-Aldrich</td>
 <td>I-6634</td>
 <td>For 1000X stock, resuspend at 5mg/ml in 5mM HCl and filter-sterilize</td>
 </tr>

 <tr>
 	<td>FBS</td>
 <td>Gibco/Invitrogen</td>
 <td>26140-079</td>
 <td>&nbsp;</td>
 </tr>

 <tr>
 	<td>ECM 830 Square-wave electroporator</td>
 <td>BTX Harvard Apparatus</td>
 <td>&nbsp;</td>
 <td>&nbsp;</td>
 </tr>

 <tr>
 	<td>Nuclepore filters</td>
 <td>Whatman</td>
 <td>110606</td>
 <td>25mm, 0.2&#x03BC;m</td>
 </tr>

 <tr>
 	<td>Tissue culture incubator</td>
 <td>&nbsp;</td>
 <td>&nbsp;</td>
 <td>37&deg;C, 5% CO2</td>
 </tr>

 <tr>
 	<td>Glass coverslips #1.5</td>
 <td>Fisher Scientific</td>
 <td>12-544E</td>
 <td>0.16mm thick</td>
 </tr>

 <tr>
 	<td>Fluorescent compound microscope equipped with camera</td>
 <td>&nbsp;</td>
 <td>&nbsp;</td>
 <td>Camera should be monochromatic (e.g., ORCA-ER camera by Hamamatsu)</td>
 </tr>

 <tr>
 	<td>EGFP/DsRed filter set for compound microscope</td>
 <td>Chroma Technology Corp.</td>
 <td>86007</td>
 <td>This filter set minimizes bleedthrough between the red and green chanenels</td>
 </tr>
 <tr>
 <td>ImageJ Software</td>
 <td>NIH</td>
 <td>&nbsp;</td>
 <td><a href="http://rsbweb.nih.gov/ij/">http://rsbweb.nih.gov/ij/</a></td>
 </tr> 
 </tbody>
</table>

quantifying the activity of cis-regulatory elements in the mouse retina by explant electroporation

Transcription factors within cellular gene networks control the spatiotemporal pattern and levels of expression of their target genes by binding to cis-regulatory elements (CREs), short (&tilde;300-600 bp) stretches of genomic DNA which can lie upstream, downstream, or within the introns of the genes they control. CREs (i.e., enhancers/promoters) typically consist of multiple clustered binding sites for both transcriptional activators and repressors1-3. They serve as logical integrators of transcriptional input giving a unitary output in the form of spatiotemporally precise and quantitatively exact promoter activity. Most studies of mammalian cis-regulation to date have relied on mouse transgenesis as a means of assaying the enhancer function of CREs4-5. This technique is time-consuming, costly and, on account of insertion site effects, largely non-quantitative. On the other hand, quantitative assays for mammalian CRE function have been developed in tissue culture systems (e.g., dual luciferase assays), but the in vivo relevance of these results is often uncertain. 
Electroporation offers an excellent alternative to traditional mouse transgenesis in that it permits both spatiotemporal and quantitative assessment of cis-regulatory activity in living mammalian tissue. This technique has been particularly useful in the analysis of cis-regulation in the central nervous system, especially in the cerebral cortex and the retina6-8. While mouse retinal electroporation, both in vivo and ex vivo, has been developed and extensively described by Matsuda and Cepko6-7,9, we have recently developed a simple approach to quantify the activity of photoreceptor-specific CREs in electroporated mouse retinas10. Given that the amount of DNA that is introduced into the retina by electroporation can vary from experiment to experiment, it is necessary to include a co-electroporated 'loading control' in all experiments. In this respect, the technique is very similar to the dual luciferase assay used to quantify promoter activity in cultured cells. 
When assaying photoreceptor cis-regulatory activity, electroporation is usually performed in newborn mice (postnatal day 0, P0) which is the time of peak rod production11-12. Once retinal cell types become post-mitotic, electroporation is much less efficient. Given the high rate of rod birth in newborn mice and the fact that rods constitute more than 70% of the cells in the adult mouse retina, the majority of cells that are electroporated at P0 are rods. For this reason, rod photoreceptors are the easiest retinal cell type to study via electroporation. The technique we describe here is primarily useful for quantifying the activity of photoreceptor CREs.

Watch this Scientific Journal Video about Quantifying the Activity of cis-Regulatory Elements in the Mouse Retina by Explant Electroporation at JoVE.com

Quantifying the Activity of cis-Regulatory Elements in the Mouse Retina by Explant Electroporation

This protocol describes a simple and inexpensive way to quantify the activity of cis-regulatory elements (i.e., enhancer/promoters) in living mouse retinas via explant electroporation. DNA preparation, retinal dissection, electroporation, retinal explant culture, and post-fixation analysis and quantification are described.

Quantifying the Activity of cis-Regulatory Elements in the Mouse Retina by Explant Electroporation

Transcription factors within cellular gene networks control the spatiotemporal pattern and levels of expression of their target genes by binding to ...

quantifying-activity-cis-regulatory-elements-mouse-retina-explant

Research

JoVE Journal

Neuroscience

14.4K 조회수.  Washington University School of Medicine. 이 프로토콜은 explant의 electroporation을 통해 마우스 망막 생활 CIS - 규제 요소의 활동 (즉, 확장기 / 발기인)를 수치 간단하고 저렴한 방법을 설명합니다. DNA의 준비, 망막 절개, electroporation, 망막 explant의 문화, 포스트 고정 분석과 부량이 설명되어 있습니다.

비디오: Quantifying the Activity of cis-Regulatory Elements in the Mouse Retina by Explant Electroporation

Transfection of Mouse Retinal Ganglion Cells by in vivo Electroporation

The targeting and refinement of RGC projections to the midbrain is a popular and powerful model system for studying how precise patterns of neural connectivity form during development. In mice, retinofugal projections are arranged in a topographic manner and form eye-specific layers in the Lateral Geniculate Nucleus (dLGN) of the thalamus and the Superior Colliculus (SC). The development of these precise patterns of retinofugal projections has typically been studied by labeling populations of RGCs with fluorescent dyes and tracers, such as horseradish peroxidase1-4. However, these methods are too coarse to provide insight into developmental changes in individual RGC axonal arbor morphology that are the basis of retinotopic map formation. They also do not allow for the genetic manipulation of RGCs.
Recently, electroporation has become an effective method for providing precise spatial and temporal control for delivery of charged molecules into the retina5-11. Current retinal electroporation protocols do not allow for genetic manipulation and tracing of retinofugal projections of a single or small cluster of RGCs in postnatal mice. It has been argued that postnatal in vivo electroporation is not a viable method for transfecting RGCs since the labeling efficiency is extremely low and hence requires targeting at embryonic ages when RGC progenitors are undergoing differentiation and proliferation6. 
In this video we describe an in vivo electroporation protocol for targeted delivery of genes, shRNA, and fluorescent dextrans to murine RGCs postnatally. This technique provides a cost effective, fast and relatively easy platform for efficient screening of candidate genes involved in several aspects of neural development including axon retraction, branching, lamination, regeneration and synapse formation at various stages of circuit development. In summary we describe here a valuable tool which will provide further insights into the molecular mechanisms underlying sensory map development.

Transfection of Mouse Retinal Ganglion Cells by in vivo Electroporation

We demonstrate an in vivo electroporation protocol for transfecting single or small clusters of retinal ganglion cells (RGCs) and other retinal cell types in postnatal mice over a wide range of ages. The ability to label and genetically manipulate postnatal RGCs in vivo is a powerful tool for developmental studies.

The targeting and refinement of RGC projections to the midbrain is a popular and powerful model system for studying how precise patterns of neural ...

In vivo Electroporation of Developing Mouse Retina

The functional characterization of genes expressed during mammalian retinal development remains a significant challenge. Gene targeting to generate constitutive or conditional loss of function knockouts remains cost and labor intensive, as well as time consuming. Adding to these challenges, retina expressed genes may have essential roles outside the retina leading to unintended confounds when using a knockout approach. Furthermore, the ability to ectopically express a gene in a gain of function experiment can be extremely valuable when attempting to identify a role in cell fate specification and/or terminal differentiation.
We present a method for the rapid and efficient incorporation of DNA plasmids into the neonatal mouse retina by electroporation. The application of short electrical impulses above a certain field strength results in a transient increase in plasma membrane permeability, facilitating the transfer of material across the membrane 1,2,3,4. Groundbreaking work demonstrated that electroporation could be utilized as a method of gene transfer into mammalian cells by inducing the formation of hydrophilic plasma membrane pores allowing the passage of highly charged DNA through the lipid bilayer 5. Continuous technical development has resulted in the viability of electroporation as a method for in vivo gene transfer in multiple mouse tissues including the retina, the method for which is described herein 6, 7, 8, 9, 10. 
DNA solution is injected into the subretinal space so that DNA is placed between the retinal pigmented epithelium and retina of the neonatal (P0) mouse and electrical pulses are applied using a tweezer electrode. The lateral placement of the eyes in the mouse allows for the easy orientation of the tweezer electrode to the necessary negative pole-DNA-retina-positive pole alignment. Extensive incorporation and expression of transferred genes can be identified by postnatal day 2 (P2). Due to the lack of significant lateral migration of cells in the retina, electroporated and non-electroporated regions are generated. Non-electroporated regions may serve as internal histological controls where appropriate. 
Retinal electroporation can be used to express a gene under a ubiquitous promoter, such as CAG, or to disrupt gene function using shRNA constructs or Cre-recombinase. More targeted expression can be achieved by designing constructs with cell specific gene promoters. Visualization of electroporated cells is achieved using bicistronic constructs expressing GFP or by co-electroporating a GFP expression construct. Furthermore, multiple constructs may be electroporated for the study of combinatorial gene effects or simultaneous gain and loss of function of different genes. Retinal electroporation may also be utilized for the analysis of genomic cis-regulatory elements by generating appropriate expression constructs and deletion mutants. Such experiments can be used to identify cis-regulatory regions sufficient or required for cell specific gene expression 11. Potential experiments are limited only by construct availability.

In vivo Electroporation of Developing Mouse Retina

A method for the incorporation of plasmid DNA into murine retinal cells for the purpose of performing either gain- or loss of function studies in vivo is presented. This method capitalizes on the transient increase in permeability of cell plasma membranes induced by the application of an external electrical field.

The functional characterization of genes expressed during mammalian retinal development remains a significant challenge.  Gene targeting to generate ...

Gene Transfer to the Developing Mouse Inner Ear by In Vivo Electroporation

The mammalian inner ear has 6 distinct sensory epithelia: 3 cristae in the ampullae of the semicircular canals; maculae in the utricle and saccule; and the organ of Corti in the coiled cochlea. The cristae and maculae contain vestibular hair cells that transduce mechanical stimuli to subserve the special sense of balance, while auditory hair cells in the organ of Corti are the primary transducers for hearing 1. Cell fate specification in these sensory epithelia and morphogenesis of the semicircular canals and cochlea take place during the second week of gestation in the mouse and are largely completed before birth 2,3. Developmental studies of the mouse inner ear are routinely conducted by harvesting transgenic embryos at different embryonic or postnatal stages to gain insight into the molecular basis of cellular and/or morphological phenotypes 4,5. We hypothesize that gene transfer to the developing mouse inner ear in utero in the context of gain- and loss-of-function studies represents a complimentary approach to traditional mouse transgenesis for the interrogation of the genetic mechanisms underlying mammalian inner ear development6.
The experimental paradigm to conduct gene misexpression studies in the developing mouse inner ear demonstrated here resolves into three general steps: 1) ventral laparotomy; 2) transuterine microinjection; and 3) in vivo electroporation. Ventral laparotomy is a mouse survival surgical technique that permits externalization of the uterus to gain experimental access to the implanted embryos7. Transuterine microinjection is the use of beveled, glass capillary micropipettes to introduce expression plasmid into the lumen of the otic vesicle or otocyst. In vivo electroporation is the application of square wave, direct current pulses to drive expression plasmid into progenitor cells8-10. 
We previously described this electroporation-based gene transfer technique and included detailed notes on each step of the protocol11. Mouse experimental embryological techniques can be difficult to learn from prose and still images alone. In the present work, we demonstrate the 3 steps in the gene transfer procedure. Most critically, we deploy digital video microscopy to show precisely how to: 1) identify embryo orientation in utero; 2) reorient embryos for targeting injections to the otocyst; 3) microinject DNA mixed with tracer dye solution into the otocyst at embryonic days 11.5 and 12.5; 4) electroporate the injected otocyst; and 5) label electroporated embryos for postnatal selection at birth. We provide representative examples of successfully transfected inner ears; a pictorial guide to the most common causes of otocyst mistargeting; discuss how to avoid common methodological errors; and present guidelines for writing an in utero gene transfer animal care protocol.

Gene Transfer to the Developing Mouse Inner Ear by In Vivo Electroporation

The mouse inner ear is a placode-derived sensory organ whose developmental program is elaborated during gestation. We define an in utero gene transfer technique consisting of three steps: mouse ventral laparotomy, transuterine microinjection, and in vivo electroporation. We use digital video microscopy to demonstrate the critical experimental embryological techniques.

The mammalian inner ear has 6 distinct sensory epithelia: 3 cristae in the ampullae of the semicircular canals; maculae in the utricle and saccule; and ...

Genetic Manipulation of the Mouse Developing Hypothalamus through In utero Electroporation

Genetic modification of specific regions of the developing mammalian brain is a very powerful experimental approach. However, generating novel mouse mutants is often frustratingly slow. It has been shown that access to the mouse brain developing in utero with reasonable post-operatory survival is possible. Still, results with this procedure have been reported almost exclusively for the most superficial and easily accessible part of the developing brain, i.e. the cortex. The thalamus, a narrower and more medial region, has proven more difficult to target. Transfection into deeper nuclei, especially those of the hypothalamus, is perhaps the most challenging and therefore very few results have been reported. Here we demonstrate a procedure to target the entire hypothalamic neuroepithelium or part of it (hypothalamic regions) for transfection through electroporation. The keys to our approach are longer narcosis times, injection in the third ventricle, and appropriate kind and positioning of the electrodes. Additionally, we show results of targeting and subsequent histological analysis of the most recessed hypothalamic nucleus, the mammillary body.

Genetic Manipulation of the Mouse Developing Hypothalamus through In utero Electroporation

Despite the functional and medical importance of the hypothalamus, in utero genetic manipulation of its development has rarely been attempted. We show a detailed procedure for in utero electroporation into the mouse hypothalamus and show representative results of total and partial (regional) hypothalamic transfection.

Genetic  modification of specific regions of the developing mammalian brain is a very  powerful experimental approach. However, generating novel mouse ...

Assessment of Vascular Regeneration in the CNS Using the Mouse Retina

The rodent retina is perhaps the most accessible mammalian system in which to investigate neurovascular interplay within the central nervous system (CNS). It is increasingly being recognized that several neurodegenerative diseases such as Alzheimer&#8217;s, multiple sclerosis, and amyotrophic lateral sclerosis present elements of vascular compromise. In addition, the most prominent causes of blindness in pediatric and working age populations (retinopathy of prematurity and diabetic retinopathy, respectively) are characterized by vascular degeneration and failure of physiological vascular regrowth. The aim of this technical paper is to provide a detailed protocol to study CNS vascular regeneration in the retina. The method can be employed to elucidate molecular mechanisms that lead to failure of vascular growth after ischemic injury. In addition, potential therapeutic modalities to accelerate and restore healthy vascular plexuses can be explored. Findings obtained using the described approach may provide therapeutic avenues for ischemic retinopathies such as that of diabetes or prematurity and possibly benefit other vascular disorders of the CNS.

The rodent retina has long been recognized as an accessible window to the brain. In this technical paper we provide a protocol that employs the mouse model of oxygen-induced retinopathy to study the mechanisms that lead to failure of vascular regeneration within the central nervous system after ischemic injury. The described system can also be harnessed to explore strategies to promote regrowth of functional blood vessels within the retina and CNS.

The rodent retina is perhaps the most accessible mammalian system in which to investigate neurovascular interplay within the central nervous system (CNS). ...

Telomerase Activity in the Various Regions of Mouse Brain: Non-Radioactive Telomerase Repeat Amplification Protocol (TRAP) Assay

Telomerase, a ribonucleoprotein, is responsible for maintaining the telomere length and therefore promoting genomic integrity, proliferation, and lifespan. In addition, telomerase protects the mitochondria from oxidative stress and confers resistance to apoptosis, suggesting its possible importance for the surviving of non-mitotic, highly active cells such as neurons. We previously demonstrated the ability of novel telomerase activators to increase telomerase activity and expression in the various mouse brain regions and to protect motor neurons cells from oxidative stress. These results strengthen the notion that telomerase is involved in the protection of neurons from various lesions. To underline the role of telomerase in the brain, we here compare the activity of telomerase in male and female mouse brain and its dependence on age. TRAP assay is a standard method for detecting telomerase activity in various tissues or cell lines. Here we demonstrate the analysis of telomerase activity in three regions of the mouse brain by non-denaturing protein extraction using CHAPS lysis buffer followed by modification of the standard TRAP assay.
In this 2-step assay, endogenous telomerase elongates a specific telomerase substrate (TS primer) by adding TTAGGG 6 bp repeats (telomerase reaction). The telomerase reaction products are amplified by PCR reaction creating a DNA ladder of 6 bp increments. The analysis of the DNA ladder is made by 4.5% high resolution agarose gel electrophoresis followed by staining with highly sensitive nucleic acid stain.
Compared to the traditional TRAP assay that utilize 32P labeled radioactive dCTP&#39;s for DNA detection and polyacrylamide gel electrophoresis for resolving the DNA ladder, this protocol offers a non-toxic time saving TRAP assay for evaluating telomerase activity in the mouse brain, demonstrating the ability to detect differences in telomerase activity in the various female and male mouse brain region.

Telomerase Activity in the Various Regions of Mouse Brain: Non-Radioactive Telomerase Repeat Amplification Protocol TRAP Assay

Telomerase is expressed in the neonatal brain and also in distinct regions of the adult brain. We present a non-toxic time saving TRAP assay for the analysis of telomerase activity in various regions of the mouse brain and detection of differences in telomerase activity between male and female mouse brains.

Telomerase, a ribonucleoprotein, is responsible for maintaining the telomere length and therefore promoting genomic integrity, proliferation, and ...

Imaging Ca2+ Dynamics in Cone Photoreceptor Axon Terminals of the Mouse Retina

Retinal cone photoreceptors (cones) serve daylight vision and are the basis of color discrimination. They are subject to degeneration, often leading to blindness in many retinal diseases. Calcium (Ca2+), a key second messenger in photoreceptor signaling and metabolism, has been proposed to be indirectly linked with photoreceptor degeneration in various animal models. Systematically studying these aspects of cone physiology and pathophysiology has been hampered by the difficulties of electrically recording from these small cells, in particular in the mouse where the retina is dominated by rod photoreceptors. To circumvent this issue, we established a two-photon Ca2+ imaging protocol using a transgenic mouse line that expresses the genetically encoded Ca2+ biosensor TN-XL exclusively in cones and can be crossbred with mouse models for photoreceptor degeneration. The protocol described here involves preparing vertical sections (&#8220;slices&#8221;) of retinas from mice and optical imaging of light stimulus-evoked changes in cone Ca2+ level. The protocol also allows &#8220;in-slice measurement&#8221; of absolute Ca2+ concentrations; as the recordings can be followed by calibration. This protocol enables studies into functional cone properties and is expected to contribute to the understanding of cone Ca2+ signaling as well as the potential involvement of Ca2+ in photoreceptor death and retinal degeneration.

Imaging Ca2+ Dynamics in Cone Photoreceptor Axon Terminals of the Mouse Retina

We describe a protocol to monitor Ca2+ dynamics in the axon terminals of cone photoreceptors using an ex-vivo&#160;slice preparation of the mouse retina. This protocol allows comprehensive studies of cone Ca2+ signaling in an important mammalian model system, the mouse.

Retinal cone photoreceptors (cones) serve daylight vision and are the basis of color discrimination. They are subject to degeneration, often leading to ...

In Vivo Gene Transfer to Schwann Cells in the Rodent Sciatic Nerve by Electroporation

The formation of the myelin sheath by Schwann cells (SCs) is essential for rapid conduction of nerve impulses along axons in the peripheral nervous system. SC-selective genetic manipulation in living animals is a powerful technique for studying the molecular and cellular mechanisms of SC myelination and demyelination in vivo. While knockout/knockin and transgenic mice are powerful tools for studying SC biology, these methods are costly and time consuming. Viral vector-mediated transgene introduction into the sciatic nerve is a simpler and less laborious method. However, viral methods have limitations, such as toxicity, transgene size constraints, and infectivity restricted to certain developmental stages. Here, we describe a new method that allows selective transfection of myelinating SCs in the rodent sciatic nerve using electroporation. By applying electric pulses to the sciatic nerve at the site of plasmid DNA injection, genes of interest can be easily silenced or overexpressed in SCs in both neonatal and more mature animals. Furthermore, this in vivo electroporation method allows for highly efficient simultaneous expression of multiple transgenes. Our novel technique should enable researchers to efficiently manipulate SC gene expression, and facilitate studies on SC development and function.

In Vivo Gene Transfer to Schwann Cells in the Rodent Sciatic Nerve by Electroporation

Here, we present an in vivo technique for gene transfer to Schwann cells (SCs) in the rodent sciatic nerve. This simple technique is useful for investigating signaling mechanisms involved in the development and maintenance of myelinating SCs.

The formation of the myelin sheath by Schwann cells (SCs) is essential for rapid conduction of nerve impulses along axons in the peripheral nervous ...

Investigating Mammalian Axon Regeneration: In Vivo Electroporation of Adult Mouse Dorsal Root Ganglion

Electroporation is an essential non-viral gene transfection approach to introduce DNA plasmids or small RNA molecules into cells. A sensory neuron in the dorsal root ganglion (DRGs) extends a single axon with two branches. One branch goes to the peripheral nerve (peripheral branch), and the other branch enters the spinal cord through the dorsal root (central branch). After the neural injury, the peripheral branch regenerates robustly whereas the central branch does not regenerate. Due to the high regenerative capacity, sensory axon regeneration has been widely used as a model system to study mammalian axon regeneration in both the peripheral nervous system (PNS) and the central nervous system (CNS). Here, we describe a previously established approach protocol to manipulate gene expression in mature sensory neurons in vivo via electroporation. Based on transfection with plasmids or small RNA oligos (siRNAs or microRNAs), the approach allows for both loss- and gain-of-function experiments to study the roles of genes-of-interests or microRNAs in regulation of axon regeneration in vivo. In addition, the manipulation of gene expression in vivo can be controlled both spatially and temporally within a relatively short time course. This model system provides a unique tool to investigate the molecular mechanisms by which mammalian axon regeneration is regulated in vivo.

Investigating Mammalian Axon Regeneration: In Vivo Electroporation of Adult Mouse Dorsal Root Ganglion

Electroporation is an effective approach to deliver genes of interests into cells. By applying this approach in vivo on the neurons of adult mouse dorsal root ganglion (DRG), we describe a model to study axon regeneration in vivo.

Electroporation is an essential non-viral gene transfection approach to introduce DNA plasmids or small RNA molecules into cells. A sensory neuron in the ...

Quantifying the Heterogeneous Distribution of a Synaptic Protein in the Mouse Brain Using Immunofluorescence

The presence, absence, or levels of specific synaptic proteins can severely influence synaptic transmission. In addition to elucidating the function of a protein, it is vital to also determine its distribution. Here, we describe a protocol employing immunofluorescence, confocal microscopy, and computer-based analysis to determine the distribution of the synaptic protein Mover (also called TPRGL or SVAP30). We compare the distribution of Mover to that of the synaptic vesicle protein synaptophysin, thereby determining the distribution of Mover in a quantitative manner relative to the abundance of synaptic vesicles. Notably, this method could potentially be implemented to allow for comparison of the distribution of proteins using different antibodies or microscopes or across different studies. Our method circumvents the inherent variability of immunofluorescent stainings by yielding a ratio rather than absolute fluorescence levels. Additionally, the method we describe enables the researcher to analyze the distribution of a protein on different levels: from whole brain slices to brain regions to different subregions in one brain area, such as the different layers of the hippocampus or sensory cortices. Mover is a vertebrate-specific protein that is associated with synaptic vesicles. With this method, we show that Mover is heterogeneously distributed across brain areas, with high levels in the ventral pallidum, the septal nuclei, and the amygdala, and also within single brain areas, such as the different layers of the hippocampus.

Here, we describe a quantitative approach to determining the distribution of a synaptic protein relative to a marker protein using immunofluorescence staining, confocal microscopy, and computer-based analysis.

The presence, absence, or levels of specific synaptic proteins can severely influence synaptic transmission. In addition to elucidating the function of a ...