Name: in utero and ex vivo electroporation for gene expression in mouse retinal ganglion cells
Uploaded: 2009-09-24T15:00:00
Duration: 25 min 6 s
Description: Watch this Scientific Journal Video about In utero and ex vivo Electroporation for Gene Expression in Mouse Retinal Ganglion Cells at JoVE.com

We thank Dr. Richard Vallee and Brikha Shrestha for help with the in utero and ex vivo retinal electroporation techniques, respectively, and Dr. T. Sakurai for comments on this protocol. This work was supported by National Institutes of Health Grants F31 NS051008 (T.J.P), the Fondation pour la Recherche Medicale, and the Human Frontier Science Program (A.R.) and R01 EY12736 (C.M.).

Part 1: Setting up for in utero and ex vivo retinal electroporations
1. For both in utero and ex vivo retinal electroporations
<ul>
 <li>Use an electrode puller to prepare fine-tipped micropipettes and break the tip (or bevel the tip) to make an angled point.</li>
 <li>Prepare DNA solutions to desired concentrations and add a small amount of Fast Green Dye (0.05%) to visualize injections. I recommend 5 &mu;l DNA solution per mother for in utero retinal electroporations and 1 &mu;l DNA solution per embryonic head for ex vivo retinal electroporations. Include a GFP expression construct (or other fluorescent protein) to visualize transfected neurons.</li>
 <li>Sterilize surgical instruments via autoclave.</li>
 <li>Prepare the ketamine-xylazine anesthetic mix with the following ratio (making ~0.2 ml for each pregnant female): a 1.0:0.2:4.6 volumetric ratio of 100 &mu;g/ml ketamine:100 &mu;g/ml xylazine:saline. Other anesthetics may be used.</li>
</ul>
2. For in utero retinal electroporations only
<ul>
 <li>Turn on the heating pad and cover it with a diaper pad.</li>
 <li>Prepare the antibiotic-antimycotic solution (1:100) in phosphate buffer saline (PBS) and warm on heating pad. Prepare ~2 ml for each dam.</li>
 <li>Place sterile-filtered PBS in warm water bath.</li>
</ul>
3. For ex vivo retinal electroporations only
 <ul>
 <li>Prepare oxygenated serum free medium (SFM) for retina incubation: Bubble 25 ml of DMEM/F12 with 95% oxygen/5% carbon dioxide for &ge; 2 hours. To the 25 ml oxygenated medium, add 0.25 g bovine serum albumin (BSA), 250 &mu;l ITS supplement and 50 &mu;l Penicillin-streptomycin solution. Mix the solution to dissolve the BSA, then sterile filter the solution. This solution should be made fresh for each electroporation experiment.</li>
 <li>Prepare SFM + 0.4% methylcellulose medium for retinal explant incubation: Autoclave 0.20 g of methylcellulose in a 150 ml bottle with a stir magnet. Prepare 50 ml of SFM (as above, without bubbling) and add to autoclaved methylcellulose bottle. Mix this solution at 4&deg;C overnight. Sterile filter this solution the following day and store at 4&deg;C, this solution can be kept for ~1 month.</li>
 <li>On the day of retinal explant harvesting (Part 5), prepare dishes for retinal explant plating: Coat glass bottom culture dishes with 250 &mu;l of poly-L-ornithine for &ge; 2 hours at 37&deg;C. (Alternatively, dishes can be coated with poly-L-ornithine the previous day and incubated overnight at 4&deg;C.) Wash dishes several times with PBS, then coat dishes with 200 &mu;l of 10 &mu;g/ml laminin in DMEM/F12 for &ge; 2 hours at 37&deg;C. Wash dishes several times with PBS at 37&deg;C and leave on PBS until explants are ready for plating.</li>
 <li>For border assays, coat dishes with poly-ornithine as above, then coat half of the dish with protein of interest along with a fluorescent marker to visualize the border (such as Alexa Fluor 555 conjugated to BSA, 1:800) for 2 hours at 37&deg;C. Wash with PBS, then add laminin as described above.</li>
 </ul>
Part 2A: In utero retinal electroporation Injecting and electroporating DNA into E14.5 murine retinae in vivo
<ol>
<li>Inject 0.15 ml of the anesthetic mix intraperitoneal (i.p.) into an E13.5-E14.5 stage dam. It should take 3-7 minutes until mother has reached a surgical plane of anesthesia. A fully anesthetized mouse should not respond to pinching of the hindpaw.</li>
<li>During this time, cut a small piece of parafilm and pipet ~5 ul of DNA solution onto the parafilm. Bend a plunger (metal wire) at a 90&deg; degree angle and insert it into a pulled micropippette. Using the plunger, aspirate the DNA solution into the micropipette (using a dissecting microscope can aid this process).</li>
<li>Once the mother has reached a surgical plane of anesthesia, use an electric razor to shave a ~2 inch region of the mother s abdomen, then place her dorsal side down on the heating pad. Prep the shaved area by wiping with an alcohol pad followed by an iodine pad a minimum of two times. Place a drop of ophthalmic Vet ointment on each eye to prevent corneal ulceration of the eyes while the mother is under general anesthesia. <img src="/files/ftp_upload/1333/1333_v2fig1.jpg" alt="Figure 1" /></li>
<li>With the forceps, grasp the skin and use the scissors to make a vertical incision along the midline (~1 inch long) through the skin and muscle of the abdomen, exposing the peritoneum. Lift the peritoneum with the forceps and make a similar vertical incision through this membrane to expose the abdominal cavity (Figure 1). Then drape the surgical field with gauze pads to prevent the abdominal contents from contacting and being contaminated by the hair of the animal.</li>
<li>Pull back the skin and peritoneum with the forceps and use the dissecting spatula to extract an embryo through the incision (choose the most accessible embryo). Place the spatula between 2 embryos and gently pull the embryonic chain out of the abdominal cavity. You can pull out the embryonic chain with your fingers. 
 *From this point forward, keep the embryos hydrated with sterile PBS*</li>
<li>Keep the mother on the heating pad, place her under the dissecting microscope and massage an embryo so that the retina is facing up. I recommend starting with either the most medial or lateral embryo, making it easier to keep track of which embryos were electroporated.</li>
<li>Take the micropipette in your dominant hand and stabilize the embryo with your other hand. With the micropipette, pierce the retina through the uterine wall and amniotic sac. Depending on the sharpness of the micropipette, it may take a fair amount of pressure to penetrate the uterine wall, but once through, the micropipette will enter the embryo and ideally the retina. It can be difficult to control and gauge the depth of the micropipette as it enters the embryo. Minimize the amount of micropipette movement once it has pierced the membrane to prevent enlarging the entrance hole, resulting in the leakage of amniotic fluid and embryo death. Avoid piercing blood vessels in the uterine wall, as this will result in bleeding and embryo death.</li>
<li>Once you are confident that the micropipette has entered the retina, you are ready to expel the DNA solution into the retina. Slowly depress the plunger and simultaneously withdraw the micropipette, ensuring that DNA is injected throughout the path of the micropipette, as the micropipette often penetrates deep to the retina. If you are in the retina, expel 0.5 &mu;l (can be measured by the 1 &mu;l tick marks on the micropipette) into the extraretinal space between the outer retinal layer and the retinal pigment epithelium (RPE). You should observe green dye filling the retina and leaks out into the amniotic fluid (Figure 1).</li>
<li>The electroporation paddles should be kept in a PBS-filled Petri dish during the surgery. Gently place the paddles on the sides of the embryo head with the positive (+) paddle on the same side as the injected eye. Apply a moderate amount of pressure to stabilize the head, but do not press too hard, as this will pop the amniotic sac and result in embryo death. When the electrodes are in place, deliver 5 40-V, 50 ms square pulses at a frequency of 60 Hz. It is not unusual for the mother to twitch due to current escaping to the skin and muscles.</li>
<li>Repeat steps 7-9 for each embryo to be injected and electroporated. I usually electroporate 4-8 embryos per mother, depending on amount of DNA, number of embryos, condition of mother, and the amount of time under anesthetic. Alternatively, multiple embryos can be injected and then electroporated, as opposed to injecting and electroporating each embryo separately.</li>
<li>After electroporating the embryos, use the forceps and spatula to replace the embryonic chain back into the abdominal cavity. Pour ~2 ml of the antibiotic-antimycotic solution into the abdominal cavity.</li>
<li>Use the hemostat and forceps to suture the peritoneum, starting with a double knot on the anterior portion of the incision and proceed with a simple continuous suture pattern to the posterior portion of the incision. Use the final loop to tie off the suture and trim excess suture.</li>
<li>To staple the incision closed, lift the skin at the anterior portion of the incision with blunt forceps, being careful to separate the skin from the peritoneum. Place the teeth of the stapler around the skin and depress the stapler. Continue stapling the incision closed to the posterior end, which usually requires 5-7 staples. Wipe the wound around the staples with an alcohol pad.</li>
<li>Let the mother recover on the heating pad (usually takes between 15-90 minutes for her to awaken) and when she begins to roll over, place her in a new cage abdominal side up.</li>
<li>To minimize pain and discomfort, mothers receive buprenorphine (0.05-0.1 mg/kg, s.c.) upon awakening, and at later time points if discomfort continues. To prevent dehydration, inject the mother with 1.0 ml saline subcutaneously (s.c.) ~2-4 hours post-surgery, and again if the evening and following day if needed. 24 hours post-surgery, the mother should regain normal behavior and be drinking and eating regularly.</li>
</ol>
Part 2B: In utero retinal electroporation Retrieval of electroporated retina at E18.5
<ol>
<li>Prepare 4% paraformaldehyde (PFA) in PBS, making ~5 ml per embryo. Perfusions can be done with a gravity-based infusion system or a syringe pump. (Alternatively, E18.5 heads can simply be immersed in 4% PFA overnight.)</li>
<li>Inject the mother with 0.2 ml of anesthetic mix i.p. and ensure that she is fully under anesthetic. Use a scissors to cut the skin and peritoneum underneath the staples to expose the abdominal cavity and uterine horns.</li>
<li>Remove an electroporated embryo and pin it ventral side up. Cut through the abdominal skin, peritoneum, diaphragm and lateral portions of the ribcage to expose the heart. Pierce the left ventricle with the butterfly needle attached to the perfusion setup, and cut the right atrium with a microscissors. Start the perfusion and allow PFA to flow for ~60 seconds; blood should exit the right atrium and the embryo should become pale if the perfusion is successful. Decapitate the embryo and collect head in 4% PFA in 15-ml conical vial. Repeat this procedure for all electroporated embryos. Keep heads in 4% PFA overnight at 4&deg;C, and then wash in PBS for several days. Euthanize mother via cervical dislocation when all embryos are collected.</li>
</ol>
Removing retina
<ol start="4">
 <li>After PBS washes, pin the head in a dissection dish with the retina facing up and immerse in PBS. Remove skin from around the retina to reveal the RPE. Using a 26G1/2 needle, puncture the ventral retina at the lens-RPE junction. Insert one edge of a microscissors into this hole and make a vertical incision through the ventral retina to the optic disc. This will allow you to orient the retina when you remove it.</li>
 <li>Remove the RPE with fine-point forceps. Grab the RPE at the ventral incision with one forceps and use other forceps to peel the RPE away from the retina, specifically at the RPE-lens junction where the RPE is firmly attached. After removing the RPE, grasp the lens with fine forceps and pull up and away to remove the lens. Remove retina by placing forceps around optic nerve head and pinch off retina. Transfer retina to a PBS-filled well, keeping track of which retinae came from which heads.</li>
 <li>Repeat steps 4-5 for the contralateral retina, and for all other electroporated embryos. The non-injected retina serves as a control for immunostaining.</li>
 <li>Use a fluorescent dissecting microscope to scan all retinae for GFP+ cells (green retinae). For all GFP+ retinae, the electroporation was successful in this embryo, and these retinae and corresponding brain (with the visual system intact) can be processed for further analysis (i.e. sectioning and immunostaining) (Figure 1).</li>
 </ol>
Part 3a: Ex vivo retinal electroporation Injecting and electroporating DNA into E14.5 murine retinae in vitro
<ol>
 <li>Anesthetize an E13.5-E14.5 stage mother with 0.2 ml of the anesthetic mix and ensure that she is fully under anesthetic. Use scissors to cut through skin and abdominal cavity to expose embryos. Lift up embryonic chain with forceps and cut through connective tissue to fully remove entire embryonic chain from the mother. Place embryonic chain in a Petri dish with DMEM/F12 on ice. Euthanize mother via cervical dislocation.</li>
 <li>Cut through the entire uterine wall with a microscissors. Then use forceps to remove each embryo from the amniotic sac and pinch off placenta. Decapitate each embryo and collect heads in another Petri dish with DMEM/F12 on ice.</li>
 <li>Using mouth pipet or plunger pipet, fill micropipette with desired amount of DNA solution (see Part 1). </li>
 <li>In this protocol, DNA will be injected into the peripheral dorsal retina. One can target other retinal regions by adjusting the DNA injection site and the positioning of the electrode paddles. </li>
 <li>Transfer one head to dissecting dish with PBS and pin the head dorsal side up. Use forceps to remove skin above retina.</li>
 <li>Holding the micropipette ~10&deg; from horizontal position (essentially parallel to the curvature of the dorsal retina), push micropipette through the RPE so that the tip is between the RPE and outer retinal layer. Expel ~0.1-0.4 &mu;l DNA solution into this space; green liquid should be observable filling entire space and leaking out through hole in RPE. Repeat injection for the contralateral retina.</li>
 <li>Carefully but quickly remove the pins and (keeping head immersed in PBS) place electrode paddles around the head, with the positive (+) paddle on the ventral side of the head. Deliver 4 40-V, 50 ms pulses at 60 Hz, then place the head back into a dish with DMEM/F12 on ice (Figure 2). <img src="/files/ftp_upload/1333/1333_v2fig2.jpg" alt="Figure 2" /></li>
 <li>Repeat steps 5-7 for all heads and for all DNA solutions. After completing electroporations, add ~1.5 ml of oxygenated SFM solution to a 4-well dish and keep on ice (one well for each different DNA solution injected).</li>
 <li>Transfer one head to a dissecting dish with DMEM/F12, with one retina facing up. On ventral retina (or side opposite target region), use fine forceps to pinch and tear a hole in the RPE. Insert one end of the forceps through this hole to peel the RPE away from the retina, specifically at the lens-retina junction. Do not cut or damage the retina because this will cause retina to collapse onto itself during incubation. After removing RPE, use forceps to pop out retina by pinching optic nerve at the base of the retina (leave the lens intact). Transfer retinae to oxygenated SFM. Flip head over and repeat for contralateral eye.</li>
 <li>Complete step 9 for all electroporated heads. Incubate retina in oxygenated SFM at 37&deg;C for 40-48 hours.</li>
</ol>
Part 3b: Ex vivo retinal electroporation Harvesting and plating of GFP+ retinal explants (2 days post-electroporation)
<ol>
 <li>Transfer all retinae from one well of oxygenated SFM into a Petri dish lid with DMEM/F12. Select one retina and use fluorescent dissecting scope to visualize GFP+ (green) portion of retina. </li>
 <li>Using a microscalpel and forceps, dissect and discard the non-GFP+ portion of retina so that only a GFP+ chunk of retina remains. It is helpful to continually switch between fluorescent and regular light throughout the dissection to specifically select the GFP+ retina. Transfer the GFP+ retina to a well with DMEM/F12.</li>
 <li>Repeat step 2 for all retina from one well. For each DNA condition (each different well), use a new Petri dish and DMEM/F12 medium.</li>
 <li>These GFP+ retinae must now be dissected into smaller retinal explants for plating. Under a dissecting scope, cut large piece of GFP+ retina into smaller explants with a microscalpel. Explants should be rectangular, ~200-300 &mu;m in length and width. Collect all explants in a well with DMEM/F12. One GFP+ chunk of retina should produce ~4-10 GFP+ retinal explants depending on tissue size. Repeat this step for all GFP+ retinal regions that were harvested.</li>
 <li>After preparing all GFP+ retinal explants, add 250 &mu;l of SFM + 0.4% methylcellulose (kept at 4&deg;C) to laminin-coated culture dishes (see Part 1). Transfer 4-8 GFP+ explants to the culture dish and use the forceps to gently arrange the explants into a polygon, with explants located halfway between the center and the edge of the dish.</li>
 <li>Determine which side of the explant is the RGC layer. Sometimes RPE crystals remain attached to the outer retinal layer, indicating that the opposite side is the RGC layer. In addition, the RGC side usually has some cellular debris that can aid in the proper explant orientation.</li>
 <li>With the RGC layer down, use forceps to firmly depress the center of the explant so that it will adhere to the glass coverslip. An indentation in the explant from the forceps may be visible; this is normal.</li>
 <li>After plating all explants on a culture dish, transfer to a 37&deg;C incubator. Retinal axons are first observed 4-6 hours after plating and explants remain healthy for several days, but significant overgrowth can occur after 24 hours. Explants can be used for numerous in vitro assays, fixed with 4% PFA for 30 minutes and immunostained (Figure 2).</li>
</ol>
Border Assay
<ol start="8">
 <li>Alternatively, transfer 4 GFP+ explants to a dish that has been prepared for the border assay (see Part 1). Arrange the explants in a vertical line in the center of the dish, approximating the location of the border. </li>
 <li>Under a fluorescent dissecting microscope, plate the explants so that they are ~50-150 mm from the fluorescent border. Incubate dishes at 37&deg;C for 24 hours, allowing ample time for RGC axons to reach border substrate. Explants can be fixed in 4% PFA for 30 minutes, followed by immunostaining (Figure 2).</li>
</ol>
Representative results
<img src="/files/ftp_upload/1333/1333_v2fig3.jpg" alt="Figure 3" />

<ol>
	<li>Chalupa, L. M., Williams, R. W. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=.">.</a> Eye Retina and the Visual Systems of the Mouse. , (2008).</li><li>Matsuda, T., Cepko, C. L. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Electroporation+and+RNA+interference+in+the+rodent+retina+in+vivo+and+in+vitro.">Electroporation and RNA interference in the rodent retina in vivo and in vitro.</a> Proc Natl Acad Sci U S A. 101, 16-22 (2004).</li><li>Ishikawa, H. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Effect+of+GDNF+gene+transfer+into+axotomized+retinal+ganglion+cells+using+in+vivo+electroporation+with+a+contact+lens-type+electrode.">Effect of GDNF gene transfer into axotomized retinal ganglion cells using in vivo electroporation with a contact lens-type electrode.</a> Gene Ther. 12, 289-298 (2005).</li><li>Donovan, S. L., Dyer, M. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Preparation+and+square+wave+electroporation+of+retinal+explant+cultures.">Preparation and square wave electroporation of retinal explant cultures.</a> Nat Protoc. 1, 2710-2718 (2006).</li><li>Garcia-Frigola, C., Carreres, M. I., Vegar, C., Herrera, E. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Gene+delivery+into+mouse+retinal+ganglion+cells+by+in+utero+electroporation.">Gene delivery into mouse retinal ganglion cells by in utero electroporation.</a> BMC Dev Biol. 7, 103-10 (2007).</li><li>Petros, T. J., Shrestha, B. R., Mason, C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Specificity+and+sufficiency+of+EphB1+in+driving+the+ipsilateral+retinal+projection.">Specificity and sufficiency of EphB1 in driving the ipsilateral retinal projection.</a> J Neurosci. 29, 3463-3474 (2009).</li></ol>

In this video, we have demonstrated electroporation techniques, both in utero and ex vivo, for gene delivery into embryonic murine RGCs. In utero retinal electroporation provides a mechanism for manipulating gene expression in RGCs and visualize their axon projections in vivo. Allowing the embryos to survive postnatally permits examination of these GFP+ RGC projections into their target, the lateral geniculate nucleus (LGN). Ex vivo retinal electroporation provides a more controlled ability to target specific retinal regions for use with in vitro assays. Combining in vivo and in vitro analyses derived from these techniques allows thorough characterization of RGC development and axon projections in a subset of RGCs in an otherwise normal background. While our demonstration focuses on RGC axon guidance at the optic chiasm, both in utero and ex vivo retinal electroporations could be beneficial for studying how perturbations in gene expression effect RGC differentiation, dendritic morphology, electrophysiological properties, synapse formation and proper targeting in termination zones such as the LGN and superior colliculus.

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<td>BTX Electro Square Porator ECM 830 (with foot paddle)</td>
<td>&nbsp;</td>
<td>Harvard Apparatus</td>
<td>450052</td>
<td>&nbsp;</td>
<tr>
<td>Round tweezer electrodes</td>
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<td>Nepa Gene</td>
<td>CUY650-5 or CUY650-7</td>
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<td>Silk Sutures</td>
<td>&nbsp;</td>
<td>Henry Schein</td>
<td>101-2636</td>
<td>&nbsp;</td>
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<td>Glass micropipettes with plungers</td>
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<td>MatTek Corporation</td>
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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.

Watch this Scientific Journal Video about In utero and ex vivo Electroporation for Gene Expression in Mouse Retinal Ganglion Cells at JoVE.com

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.

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 ...

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Biology

In Utero Intraventricular Injection and Electroporation of E16 Rat Embryos

In-utero in-vivo injection and electroporation of the embryonic rat neocortex provides a powerful tool for the manipulation of individual progenitors lining the walls of the lateral ventricle. This technique is now widely used to study the processes involved in corticogenesis by over-expressing or knocking down genes and observing the effects on cellular proliferation, migration, and differentiation. In comparison to traditional knockout strategies, in-utero electroporation provides a rapid means to manipulate a population of cells during a specific temporal window. In this video protocol, we outline the experimental methodology for preparing rats for surgery, exposing the uterine horns through laporatomy, injecting DNA into the lateral ventricles of the developing embryo, electroporating DNA into the progenitors lining the lateral wall, and caring for animals post-surgery. Our laboratory uses this protocol for surgeries on E15-E21 rats, however it is most commonly performed at E16 as shown in this video.

In-utero in-vivo injection and electroporation of the embryonic rat neocortex provides a powerful tool for the manipulation of individual progenitors ...

In Utero Intraventricular Injection and Electroporation of E15 Mouse Embryos

In-utero in-vivo injection and electroporation of the embryonic mouse neocortex provides a powerful tool for the manipulation of individual progenitors lining the walls of the lateral ventricle. This technique is now widely used to study the processes involved in corticogenesis by over-expressing or knocking down genes and observing the effects on cellular proliferation, migration, and differentiation. In comparison to traditional knockout strategies, in-utero electroporation provides a rapid means to manipulate a population of cells during a specific temporal window. In this video protocol we outline the experimental methodology for preparing mice for surgery, exposing the uterine horns through laporatomy, injecting DNA into the lateral ventricles of the developing embryo, electroporating DNA into the progenitors lining the lateral wall, and caring for animals post-surgery. Our laboratory uses this protocol for surgeries on E13-E16 mice, however, it is most commonly performed at E15, as shown in this video. 

In-utero in-vivo injection and electroporation of the embryonic mouse neocortex provides a powerful tool for the manipulation of individual progenitors ...

The Preparation of Primary Hematopoietic Cell Cultures From Murine Bone Marrow for Electroporation

It is becoming increasingly apparent that electroporation is the most effective way to introduce plasmid DNA or siRNA into primary cells. The Gene Pulser MXcell  electroporation system and Gene Pulser  electroporation buffer were specifically developed to transfect nucleic acids into mammalian cells and difficult-to-transfect cells, such as primary and stem cells.This video demonstrates how to establish primary hematopoietic cell cultures from murine bone marrow, and then prepare them for electroporation in the MXcell system. We begin by isolating femur and tibia. Bone marrow from both femur and tibia are then harvested and cultures are established. Cultured bone marrow cells are then transfected and analyzed.

This procedure describes how to establish primary hematopoietic cell cultures from murine bone marrow and is followed by transfection using the Gene Pulser MXCell electroporation system.

It is becoming increasingly apparent that electroporation is the most effective way to introduce plasmid DNA or siRNA into primary cells. The Gene Pulser ...

Using the Gene Pulser MXcell Electroporation System to Transfect Primary Cells with High Efficiency

It is becoming increasingly apparent that electroporation is the most effective way to introduce plasmid DNA or siRNA into primary cells. The Gene Pulser MXcell electroporation system and Gene Pulser electroporation buffer (Bio-Rad) were specifically developed to easily transfect nucleic acids into mammalian cells and difficult-to-transfect cells, such as primary and stem cells. We will demonstrate how to perform a simple experiment to quickly identify the best electroporation conditions. We will demonstrate how to run several samples through a range of electroporation conditions so that an experiment can be conducted at the same time as optimization is performed. We will also show how optimal conditions identified using 96-well electroporation plates can be used with standard electroporation cuvettes, facilitating the switch from electroporation plates to electroporation cuvettes while maintaining the same electroporation efficiency. In the video, we will also discuss some of the key factors that can lead to the success or failure of electroporation experiments.

This procedure shows how to use the Gene Pulser MXcell electroporation system to rapidly and easily identify the best electroporation conditions for mouse embryonic fibroblasts (MEFs) or other primary cells. Considerations for troubleshooting are also discussed in the associated video.

Using an Automated Cell Counter to Simplify Gene Expression Studies: siRNA Knockdown of IL-4 Dependent Gene Expression in Namalwa Cells

The use of siRNA mediated gene knockdown is continuing to be an important tool in studies of gene expression.  siRNA studies are being conducted not only to study the effects of downregulating single genes, but also to interrogate signaling pathways and other complex interaction networks.  These pathway analyses require both the use of relevant cellular models and methods that cause less perturbation to the cellular physiology.  Electroporation is increasingly being used as an effective way to introduce siRNA and other nucleic acids into difficult to transfect cell lines and primary cells without altering the signaling pathway under investigation. There are multiple critical steps to a successful siRNA experiment, and there are ways to simplify the work while improving the data quality at several experimental stages.  To help you get started with your siRNA mediated gene knockdown project, we will demonstrate how to perform a pathway study complete from collecting and counting the cells prior to electroporation through post transfection real-time PCR gene expression analysis.  The following study investigates the role of the transcriptional activator STAT6 in IL-4 dependent gene expression of CCL17 in a Burkitt lymphoma cell line (Namalwa).  The techniques demonstrated are useful for a wide range of siRNA-based experiments on both adherent and suspension cells.  We will also show how to streamline cell counting with the TC10 automated cell counter, how to electroporate multiple samples simultaneously using the MXcell electroporation system, and how to simultaneously assess RNA quality and quantity with the Experion automated electrophoresis system.

This procedure describes a quick and easy workflow to introduce siRNA into difficult to transfect cell lines and follow gene expression by real-time PCR. Use of an automated cell counter, multi-well electroporation plate, and automated electrophoresis station provide quick and reliable results without the need for expensive robotic handling.

The use of siRNA mediated gene knockdown is continuing to be an important tool in studies of gene expression.  siRNA studies are being conducted not only ...

In vitro Reconstitution of the Active T. castaneum Telomerase

Efforts to isolate the catalytic subunit of telomerase, TERT, in sufficient quantities for structural studies, have been met with limited success for more than a decade. Here, we present methods for the isolation of the recombinant Tribolium castaneum TERT (TcTERT) and the reconstitution of the active T. castaneum telomerase ribonucleoprotein (RNP) complex in vitro.
Telomerase is a specialized reverse transcriptase1 that adds short DNA repeats, called telomeres, to the 3' end of linear chromosomes2 that serve to protect them from end-to-end fusion and degradation. Following DNA replication, a short segment is lost at the end of the chromosome3 and without telomerase, cells continue dividing until eventually reaching their Hayflick Limit4. Additionally, telomerase is dormant in most somatic cells5 in adults, but is active in cancer cells6 where it promotes cell immortality7.
The minimal telomerase enzyme consists of two core components: the protein subunit (TERT), which comprises the catalytic subunit of the enzyme and an integral RNA component (TER), which contains the template TERT uses to synthesize telomeres8,9. Prior to 2008, only structures for individual telomerase domains had been solved10,11. A major breakthrough in this field came from the determination of the crystal structure of the active12, catalytic subunit of T. castaneum telomerase, TcTERT1.
Here, we present methods for producing large quantities of the active, soluble TcTERT for structural and biochemical studies, and the reconstitution of the telomerase RNP complex in vitro for telomerase activity assays. An overview of the experimental methods used is shown in Figure 1.

In vitro Reconstitution of the Active T. castaneum Telomerase

Efforts to isolate the catalytic subunit of telomerase, TERT, in sufficient quantities for structural studies, have been met with limited success for more than a decade. Here, we present methods for the isolation of the recombinant Tribolium castaneum TERT (TcTERT) and the reconstitution of the active T. castaneum telomerase ribonucleoprotein (RNP) complex in vitro.

Efforts to isolate the catalytic subunit of telomerase, TERT, in sufficient quantities for structural studies, have been met with limited success for more ...

Using SecM Arrest Sequence as a Tool to Isolate Ribosome Bound Polypeptides

Extensive research has provided ample evidences suggesting that protein folding in the cell is a co-translational process1-5. However, the exact pathway that polypeptide chain follows during co-translational folding to achieve its functional form is still an enigma. In order to understand this process and to determine the exact conformation of the co-translational folding intermediates, it is essential to develop techniques that allow the isolation of RNCs carrying nascent chains of predetermined sizes to allow their further structural analysis.
SecM (secretion monitor) is a 170 amino acid E. coli protein that regulates expression of the downstream SecA (secretion driving) ATPase in the secM-secA operon6. Nakatogawa and Ito originally found that a 17 amino acid long sequence (150-FSTPVWISQAQGIRAGP-166) in the C-terminal region of the SecM protein is sufficient and necessary to cause stalling of SecM elongation at Gly165, thereby producing peptidyl-glycyl-tRNA stably bound to the ribosomal P-site7-9. More importantly, it was found that this 17 amino acid long sequence can be fused to the C-terminus of virtually any full-length and/or truncated protein thus allowing the production of RNCs carrying nascent chains of predetermined sizes7. Thus, when fused or inserted into the target protein, SecM stalling sequence produces arrest of the polypeptide chain elongation and generates stable RNCs both in vivo in E. coli cells and in vitro in a cell-free system. Sucrose gradient centrifugation is further utilized to isolate RNCs.
The isolated RNCs can be used to analyze structural and functional features of the co-translational folding intermediates. Recently, this technique has been successfully used to gain insights into the structure of several ribosome bound nascent chains10,11. Here we describe the isolation of bovine Gamma-B Crystallin RNCs fused to SecM and generated in an in vitro translation system.

We describe here a technique that is now routinely used to isolate stably bound ribosome nascent chain complexes (RNCs). This technique takes advantage of the discovery that a 17 amino acid long SecM "arrest sequence" can halt translation elongation in a prokaryotic (E. coli) system, when inserted into (or fused to the C-terminus) of virtually any protein.

Extensive research has provided ample evidences suggesting that protein folding in the cell is a co-translational process1-5. However, the exact pathway ...

Gene Transfer into Older Chicken Embryos by ex ovo Electroporation

The chicken embryo provides an excellent model system for studying gene function and regulation during embryonic development. In ovo electroporation is a powerful method to over-express exogenous genes or down-regulate endogenous genes in vivo in chicken embryos1. Different structures such as DNA plasmids encoding genes2-4, small interfering RNA (siRNA) plasmids5, small synthetic RNA oligos6, and morpholino antisense oligonucleotides7 can be easily transfected into chicken embryos by electroporation. However, the application of in ovo electroporation is limited to embryos at early incubation stages (younger than stage HH20 - according to Hamburg and Hamilton)8 and there are some disadvantages for its application in embryos at later stages (older than stage HH22 - approximately 3.5 days of development). For example, the vitelline membrane at later stages is usually stuck to the shall membrane and opening a window in the shell causes rupture of the vessels, resulting in death of the embryos; older embryos are covered by vitelline and allantoic vessels, where it is difficult to access and manipulate the embryos; older embryos move vigorously and is difficult to control the orientation through a relatively small window in the shell.
In this protocol we demonstrate an ex ovo electroporation method for gene transfer into chicken embryos at late stages (older than stage HH22). For ex ovo electroporation, embryos are cultured in Petri dishes9 and the vitelline and allantoic vessels are widely spread. Under these conditions, the older chicken embryos are easily accessed and manipulated. Therefore, this method overcomes the disadvantages of in ovo electroporation applied to the older chicken embryos. Using this method, plasmids can be easily transfected into different parts of the older chicken embryos10-12.

Gene Transfer into Older Chicken Embryos by ex ovo Electroporation

A method of gene transfer into chicken embryos at later incubation stages (older than Hamburger and Hamilton stage (HH) 22) is described. This method overcomes disadvantages of in ovo electroporation applied to older chicken embryos and is a useful technique to study gene function and regulation at older developmental stages.

The chicken embryo provides an  excellent model system for studying gene function and regulation during  embryonic development. In ovo electroporation is ...

siRNA Screening to Identify Ubiquitin and Ubiquitin-like System Regulators of Biological Pathways in Cultured Mammalian Cells

Post-translational modification of proteins with ubiquitin and ubiquitin-like molecules (UBLs) is emerging as a dynamic cellular signaling network that regulates diverse biological pathways including the hypoxia response, proteostasis, the DNA damage response and transcription.&#160; To better understand how UBLs regulate pathways relevant to human disease, we have compiled a human siRNA &#8220;ubiquitome&#8221; library consisting of 1,186 siRNA duplex pools targeting all known and predicted components of UBL system pathways. This library can be screened against a range of cell lines expressing reporters of diverse biological pathways to determine which UBL components act as positive or negative regulators of the pathway in question.&#160; Here, we describe a protocol utilizing this library to identify ubiquitome-regulators of the HIF1A-mediated cellular response to hypoxia using a transcription-based luciferase reporter.&#160; An initial assay development stage is performed to establish suitable screening parameters of the cell line before performing the screen in three stages: primary, secondary and tertiary/deconvolution screening. &#160;The use of targeted over whole genome siRNA libraries is becoming increasingly popular as it offers the advantage of reporting only on members of the pathway with which the investigators are most interested.&#160; Despite inherent limitations of siRNA screening, in particular false-positives caused by siRNA off-target effects, the identification of genuine novel regulators of the pathways in question outweigh these shortcomings, which can be overcome by performing a series of carefully undertaken control experiments.

Here, we describe a methodology to perform a targeted siRNA &#8220;ubiquitome&#8221; screen to identify novel ubiquitin and ubiquitin-like regulators of the HIF1A-mediated cellular response to hypoxia.&#160; This can be adapted to any biological pathway where a robust read out of reporter activity is available.

Post-translational modification of proteins with ubiquitin and ubiquitin-like molecules (UBLs) is emerging as a dynamic cellular signaling network that ...

Measurement of Metabolic Rate in Drosophila using Respirometry

Metabolic disorders are a frequent problem affecting human health. Therefore, understanding the mechanisms that regulate metabolism is a crucial scientific task. Many disease causing genes in humans have a fly homologue, making Drosophila a good model to study signaling pathways involved in the development of different disorders. Additionally, the tractability of Drosophila simplifies genetic screens to aid in identifying novel therapeutic targets that may regulate metabolism. In order to perform such a screen a simple and fast method to identify changes in the metabolic state of flies is necessary. In general, carbon dioxide production is a good indicator of substrate oxidation and energy expenditure providing information about metabolic state. In this protocol we introduce a simple method to measure CO2 output from flies. This technique can potentially aid in the identification of genetic perturbations affecting metabolic rate.

Measurement of Metabolic Rate in Drosophila using Respirometry

Metabolic disorders are among one of the most common diseases in humans. The genetically tractable model organism D. melanogaster can be used to identify novel genes that regulate metabolism. This paper describes a relatively simple method which allows studying the metabolic rate in flies by measuring their CO2 production.