The cerebral cortex is a very complex and intricately organized structure. To achieve such a degree of organization, cortical projection neurons go through complex developmental processes that require their temporal generation, migration to their final destination in the cortical plate, and the establishment of short- and long-range connections^1,2. For a long time, the classical way to study corticogenesis was based on the use of knockout or knock-in murine models of genes of interest. However, this strategy, and particularly the use of conditional knockout mice, is time consuming and expensive, and sometimes presents additional problems regarding the existence of genetic redundancy or the lack of specific CRE drivers, among other issues. One of the approaches that arose to try to address those problems and that is nowadays extensively used to study cortical development is in utero electroporation^3,4. In utero electroporation is a technique used for somatic transgenesis, allowing in vivo targeting of neural stem cells and their progeny. This method can be used to label cells by the expression of fluorescent proteins^5,6, for gene manipulation in vivo (i.e., gain or loss of function assays)^7,8,9, for isolating electroporated cortices in vitro and culturing cells^8,10. Moreover, in utero electroporation permits temporal and regional control of the targeted area. This technique has numerous applications and has been widely used to study neuronal migration, stem cell division, neuronal connectivity, and other subjects^8,9,11,12.

The current manuscript describes the use of an in utero electroporation variant, termed double in utero electroporation, to analyze the interactions of cells in the cerebral cortex with different temporal and spatial origins. These studies are extremely complex to complete when employing murine models because they require the combined use of several transgenic lines. Some of the applications of the protocol described in this paper include the study of close interactions among neighboring cells, as well as interactions between distant cells through long-range projections. The method requires performing two independent in utero electroporation surgeries, separated temporally and spatially, on the same embryos to target different cell populations of interest. The advantage of this approach is the possibility of manipulating gene function in one or both types of neurons using wild type animals. In addition, these functional experiments can be combined with the expression of cytoplasmic or membrane-tagged fluorescent proteins to visualize the fine morphology of targeted cells, including dendrites and axons, and analyze possible differences in cellular interactions in comparison with a control (i.e., cells only labeled with the fluorescent protein).

The protocol delineated here is focused on the study of cellular interactions inside the neocortex, but this strategy could also be used to examine interactions with extracortical areas that can be targeted using in utero electroporation, like the subpallium or the thalamus^13,14, or cell-cell interactions in other structures, like the cerebellum¹⁵. Targeting of different areas is based on the orientation of the electrodes and on the ventricle where the DNA is injected (lateral, third, or fourth). With the strategy described here, we can label a substantial number of cells, which is useful to evaluate general changes in connectivity/innervation in functional experiments. Nevertheless, to study fine changes in connectivity, one can use modified versions of in utero electroporation to get sparser labeling and identify single cells¹⁶. In summary, double in utero electroporation is a versatile method that allows targeting temporally and spatially separated cell populations and studying their interactions in detail, either in control conditions or combined with functional experiments, considerably reducing temporal and economic costs.

The procedure herein described has been approved by the ethical committee in charge of experimentation, the animal welfare of the Universidad de Valencia and the Conselleria de Agricultura, Desarrollo Rural, Emergencia Climática y Transición Ecológica of the Comunidad Valenciana, and adheres to the guidelines of the International Council for Laboratory Animal Science (ICLAS) reviewed in the Real Decreto 53/2013 of the Spanish legislation as well as in the Directive 2010/63/EU of the European Parliament and of the Council.

NOTE: This protocol involves two different purposes: 1) the first study, referred to as “strategy A”, allows the analysis of the interactions between Cajal-Retzius cells (CR-cells) and early-born cortical projection neurons within the same brain hemisphere; 2) the second study, “strategy B”, is carried out in order to examine the innervation of the upper layer callosal projection neurons to the contralateral side of the neocortex.

1. Presurgery preparation

1. DNA preparation
   1. Transform chemically or electrocompetent E. coli DH5α cells with the plasmids of interest, plate them on LB agar plates with the appropriate antibiotic, and incubate them overnight at 37 °C.
      NOTE: All plasmids used here contain the general promoter for chicken β-actin (CAG) driving the expression of a fluorescent-reporter protein (CAG-mCherry and CAG-EGFP for strategy A and CAG-BFP and CAG-nEGFP-2A-mtdTomato for strategy B). All of them contain resistance to ampicillin (AMP).
   2. Pick individual colonies from each plasmid transformation and initiate a starter liquid culture in 2 mL of Luria broth + ampicillin (LB+AMP) in bacterial culture tubes during 3−4 h at 37 °C with vigorous shaking (200 rpm). After, set a larger bacterial culture in a 500 mL Erlenmeyer flask adding 200 mL of LB+AMP and 1 mL of the starter culture. Incubate overnight at 37 °C in the orbital shaker at 200 rpm.
   3. Use an endotoxin-free maxi-prep kit (Table of Materials) following the manufacturer’s instructions to obtain pure and concentrated plasmid DNA from the liquid cultures. Resuspend the DNA in ~50−100 µL of endotoxin-free Tris-EDTA (TE) buffer to obtain concentrations of at least 5 µg/µL.
   4. For each surgery, prepare a solution with a final volume of 10 µL containing 1 µL of fast green dye, plasmid DNA solution of interest for a final concentration of 1 µg/µL for each plasmid, and endotoxin-free TE buffer. For example, mix 1 µL of fast green dye, 2 µL of a 5 µg/µL plasmid DNA solution, and 7 µL of TE buffer.
2. Pipette pulling
   1. Pull borosilicate glass capillaries (1/0.58 mm outer/inner diameter) in a vertical micropipette puller until the tip reaches a length of 1−1.5 cm and trim it using dissecting forceps at an angle of approximately 30° under a dissecting scope.
3. Surgery room setup
   1. Put all the equipment on the operating table (i.e., tweezers, scissors, forceps, micropipettes, needle, and needle holder). Turn on the heating pad and cover it with a sterile surgical absorbent pad. Ensure that the reservoir in the machine for inhalation anesthesia is filled with isoflurane, the oxygen tank contains enough oxygen, and the system functions properly.
      NOTE: The surgery room and the resistant material must be kept as sterile as possible (i.e., all the material must be autoclaved before the surgery and surfaces must be sanitized with 70% ethanol). Platinum electrodes need to be carefully disinfected first with germicidal soap and second with 70% ethanol prior the surgery.
   2. Prepare 100 mL of 0.9% (w/v) sterile saline solution containing penicillin-streptomycin 1:100 and fill a 10 cm Petri dish. Place the plate on top of the heated pad to warm up the solution.
   3. Fill a 1 mL syringe with 150 µL of an analgesic solution (e.g., 0.1 mg/kg buprenorphine).
   4. Load the pulled pipette with 5 µL of the final plasmid DNA solution prepared in step 1.1.4. Connect the capillary to a mouth-controlled aspirator tube.

2. First in utero electroporation surgery

1. Place an E11.5 (strategy A) or E13.5 (strategy B) C57BL/6 pregnant mouse inside a closed induction chamber with 2.5% (v/v) isoflurane at 0.8 L/min and wait until it is anesthetized. Transfer the mouse to the heating pad and put its nose into a mask for constant delivery of isoflurane. Check for the absence of a pedal reflex as an indicator of proper anesthesia.
   NOTE: Embryonic age is determined based on the day when the vaginal plug is observed (E0.5).
2. Inject the pregnant female with the analgesic solution (0.1 mg/kg buprenorphine) subcutaneously. To prevent the eyes from drying during the procedure, apply one drop of eye ointment in each eye using a cotton swab.
3. Shave the mouse abdominal area with an electric razor and wash it 2x with 70% (v/v) ethanol wipes, once with iodine wipes, and one last time with an ethanol wipe.
4. Use scissors to make a 30 mm long incision through the skin in the right side of the animal and carefully separate the adjacent skin from the muscle with a blunt spatula. Afterwards, make a second incision in the abdominal wall.
5. Cover the abdomen with a piece of folded tissue paper previously disinfected with 70% ethanol containing a 40 mm long slit in its center. Carefully pull the uterus out of the abdominal cavity with ring forceps.
   NOTE: The uterus must be kept wet with the warm saline solution prepared in step 1.3.2 during the whole procedure.
6. Preload the pulled pipettes with the DNA solution prepared in step 1.1.4. Inject ~0.5 µL of the DNA solution per embryo into the lateral ventricle of the selected hemisphere using the mouth-controlled aspirator tube until the fast green dye is noticeable inside the ventricle.
7. Place forceps-type platinum electrodes laterally around the head of the injected embryo (as shown in Figure 1A and Figure 2A). Orient the electrodes to target the desired brain region. Direct the positive pole towards the medial wall to electroporate the cortical hem (strategy A) or towards the lateral cortex (strategy B) to label cells generated in that area.
   NOTE: In all cases, the heart and placenta must be avoided to ensure embryonic survival.
8. Apply the specific sequence of electric pulses with a square wave electroporator following the indications shown in Table 1 (strategy A E11.5 embryos: four pulses of 25 V and 40 ms, separated by 950 ms intervals; strategy B E13.5 embryos: five pulses of 35 V and 60 ms, with 950 ms intervals).
9. Carefully place the uterus back into the abdominal cavity with forceps, fill it with warm saline solution, and close the abdominal wall with a needle 6-0 suture. Join the two sides of the initial incision made in the skin either using a needle 6-0 suture or suture clips.
10. Maintain the animal on the heating pad and monitor it until its recovery from anesthesia. Provide an extra dose of analgesia (150 µL) in a hydrogel solution placed in its home cage.
11. 24 hours after surgery, administer an extra dose of analgesia (150 µL). Continue daily monitoring by visual inspection for possible pain and distress. Observe the animal’s behavior, test its normal hind limb reflexes, and inspect the suture for possible signs of damage due to licking or scratching of the wound.  

3. Second in utero electroporation

1. Two days after the initial surgery, repeat steps 2.1−2.3. Although the recovery of pregnant females after the surgery is very good, check that they show normal behavior and present no signs of pain or distress before performing the second surgery (E13.5 embryos for strategy A and E15.5 for strategy B).
2. Make a 30 mm long incision through the skin as in step 2.4 and a second incision at the abdominal wall, this time in the left side of the animal. Carefully expose the uterus on top of a disinfected tissue as described in step 2.5.
   NOTE: Be careful not to interfere with the incision previously made in the other side.
3. Inject around 0.5 µL per embryo of the DNA solution into the lateral brain ventricle of the hemisphere previously electroporated in the case of strategy A and in the lateral brain ventricle of the contralateral hemisphere for strategy B.
   NOTE: Use only embryos showing normal development and no signs of reabsorption.
4. Place the electrodes around the embryo’s head as described in step 2.7, directing the positive electrode towards the lateral cortex and apply the appropriate pulses following the indications shown in Table 1 (strategy A E13.5 embryos: five pulses of 35 V and 60 ms separated by 950 ms intervals; strategy B E15.5 embryos: five pulses of 50 V and 80 ms separated by 950 ms intervals).
5. Continue and finish the surgery as described in steps 2.8−2.10.

4. Tissue harvesting and sectioning

1. Strategy A
   1. Four days after the second electroporation (E17.5), perform cervical dislocation of the pregnant female and place it in a supine position.
   2. Using scissors, make a ventral incision to extract the uterine horns and with forceps place them in a Petri dish filled with 1x phosphate-buffered saline (PBS) placed on ice.
      NOTE: The low temperature makes the anesthetization of the embryos possible.
   3. Using tweezers, extract the embryos out of the amniotic sac and transfer them with forceps to a new PBS-filled Petri dish under a dissecting scope. Carefully hold the head of the embryos using forceps and conduct brain dissection with tweezers to first remove the skin over the head and then the skull. Use a spatula to pull out the exposed brains.
   4. Collect the brains with a spoon and deposit them in a 48 well plate filled with the fixative solution (4% [w/v] paraformaldehyde [PFA] in 1x PBS). Test immediately for the successful outcome of both electroporations by examining the brains using an inverted epifluorescence microscope, for example.
   5. Fixate the embryonic brains overnight at 4 °C in an orbital shaker. Wash with 1x PBS to eliminate traces of PFA. Then transfer them to PBS with antifungal preservatives (1x PBS-0.05% (w/v) sodium azide).
      CAUTION: PFA and sodium azide are cytotoxic compounds that require special precaution during their utilization.
   6. Embed the fixated brains in 4% (w/v) low melting point agarose in 1x PBS and wait ~10 min until it solidifies. Stick them to the vibratome tissue holder using cyanoacrylate glue with the olfactory bulbs facing upwards to obtain coronal sections.
   7. Initiate the vibratome and select the desired parameters: 100 µm width, 0.60 µm/s speed, and 0.60 mm of amplitude.
   8. Secure the brain inside the vibratome container, fill it with 1x PBS solution, and begin collecting coronal serial sections with the help of a brush in a 48 well plate filled with 1x PBS-0.05% (w/v) sodium azide to have a complete depiction of the brain (e.g., around seven sections per well and six wells per embryo).
   9. Mount the desired sections in microscope glass slides using a fine brush. Cover them with glass coverslips. For long-term storage add mounting medium, which prevents photobleaching and photooxidation. Observe the slides under an upright epifluorescence microscope to assess electroporation efficacy.
2. Strategy B
   1. Let the pups previously electroporated at E13.5 and E15.5 be born and wait until P15 to perform transcardial perfusion using the same fixative solution used in step 4.1.4.
   2. Just before perfusion, intraperitoneally administer a dose of 75/1 mg/kg ketamine/medetomidine. When the pedal reflex is lost, secure the mouse in a supine position and make a ventral incision following the middle line using scissors to expose both the rib cage and diaphragm.
   3. Cut the diaphragm and open the rib cage to gain access to the heart. Hold the rib cage using a hemostat and make an incision in the right atrium with fine scissors.
   4. Penetrate the left ventricle with a needle connected with a flexible tube to a peristaltic perfusion pump. Start transcardial perfusion, delivering at least 25 mL of 4% PFA at a constant flux of 5.5 mL/min (total time ~5 min).
   5. Dissect the brain of perfused animals. First, remove the skin over the head with scissors and forceps. Start cutting the skull using scissors and carefully pull off sections of the skull bones until they are completely removed. Finally, extract the brain with the help of a spatula.
   6. Transfer the brains to a 24 well plate and fix them with 4% PFA overnight at 4 °C on an orbital shaker. Stop fixation by replacing PFA with 0.05% (w/v) sodium azide in PBS.
      NOTE: As indicated in step 4.1.5, it is advisable to carry out an intermediate wash with 1x PBS.
   7. Repeat steps 4.1.6−4.1.9, changing the vibratome parameters for postnatal brains (40 µm width, 1.20 µm/s speed, and 0.5 mm of amplitude).
      NOTE: Immunohistochemistry or immunofluorescence can be carried out to detect specific cell markers or enhance the signal of fluorescent proteins used in electroporation.

5. Confocal fluorescence imaging and analysis

1. Turn on the confocal microscope, place the microscope slides containing the mounted brain sections onto the microscope slides holder and select the channels at which fluorescence images will be taken (i.e., 420−460 nm for BFP, 490−540 nm for GFP, and 570−620 nm for mCherry and tdTomato).
2. Perform sample scanning to obtain map images of each brain section at two different wavelengths for a general view of the double electroporation output. Once finished, select the 10x lens and the multi-area-Z-stack-timelapse observation mode. This will allow programming the automatic acquisition of fluorescence images at different XYZ localizations within brain sections.
3. In each of the chosen regions (XY), set proper imaging parameters (i.e., laser intensity, photomultiplier sensitivity, and a minimum resolution of 1,024 x 1,024), as well as the depth of the scanning (Z) according to the planes of the sample where fluorescence is visible.
4. Obtain low magnification (10x) images of all the chosen regions and export them from OIF to TIFF format using the microscope viewer software.
5. Change to the 60x lens, repeat step 5.3 and capture high magnification images to observe the cell-cell interactions in a more detailed manner. Export them as indicated in step 5.4.
6. Open the acquired images with any imaging software (e.g., Fiji) for further analysis.

Interactions between neighboring cells originated in distal places and at different times: Cajal-Retzius cells (CR-cells) and early migrating cortical projection neurons (strategy A)

The interaction of CR-cells and early cortical projection neurons was previously described as necessary to regulate somal translocation via nectin and cadherin adhesion molecules using a double electroporation strategy⁸. CR-cells originate from the neuroepithelium at the edges of the pallium and migrate tangentially to populate the most superficial part of the cortex, the marginal zone^17,18,19, whereas cortical projection neurons are generated in the proliferative zone of the cerebral cortex and migrate radially into the nascent cortical plate²⁰. There is a temporal difference in the generation of both types of cells. CR-cells are generated at very early embryonic stages from E10.5^21,22 and cortical projection neurons that migrate by somal translocation are born from E12.5–E13.5²³. Using double in utero electroporation, the temporal gap between the surgeries allows CR-cells, targeted at E11.5 in one of its places of origin (the cortical hem), to reach the marginal zone of the lateral neocortex including the somatosensory area in time to establish contacts with cortical projection neurons labeled at E13.5 (Figure 1A,B). The leading processes of projection neurons expressing enhanced GFP (EGFP) profusely arborize in the marginal zone of the cortex and intermingle with the processes of CR-cells that express mCherry (Figure 1C). Functional experiments have shown that perturbation of cell-cell adhesion molecules expressed by projection neurons or CR-cells affects the arborization of their processes as a consequence of altered contacts between both cell types⁸.

Long-range interactions between distal cells generated at different times: innervation of upper layer callosal projection neurons to the contralateral side of the neocortex (strategy B)

Callosal cortical projection neurons are present throughout the cerebral cortex, being more abundant in upper layers²⁴. These neurons project their axons through the corpus callosum and contact their target cells, projection neurons located across the different layers of the contralateral cortex ^25,26,27. Upper layer projection neurons are evolutionarily newer than lower layer neurons and have been greatly expanded in primates²⁸. These cells are critical for complex thought and higher associative tasks, and dysfunctions in groups of genes specifically expressed by this population of cells have been recently related with autism²⁹.

In order to study specifically the interactions of the subpopulation of callosal projection neurons located in the upper layers with their target cells distributed throughout the contralateral hemisphere, we developed a double in utero electroporation protocol. To label the target cells of upper layer callosal projection neurons we performed in utero electroporation at E13.5 using a BFP expressing plasmid (Figure 2A−C). This age was strategically chosen because it allows not only targeting of a broad cortical projection neuron population including many layer-V neurons, but also a considerable number of neurons located in upper layers (Figure 2C), basically covering all target areas of callosal projection neurons from the contralateral hemisphere. A second electroporation in the contralateral side at E15.5 targeted the upper layer callosal projection neuron subpopulation (expressing nEGFP and mtdTomato) (Figure 2A,B,D) but not the lower layer projection neurons born at earlier ages. The need of heterochronic double electroporation was therefore justified because of the different times in the generation of the projecting cells of interest and the cells innervated by them. Those upper layer targeted neurons send their axons to the contralateral hemisphere with a characteristic arborization pattern (Figure 2C). Differences in this typical axonal arborization pattern could be evaluated upon gain or loss of function experiments in target cells using this double electroporation protocol. High magnification analysis shows in detail the callosal axons innervating targeted projection neurons in the contralateral hemisphere (Figure 2E).

Figure 1: Strategy A. Double in utero electroporation strategy to study close interactions among cells with different spatial and temporal origin. (A) Schematics of the protocol used to target CR-cells expressing mCherry and cortical projection neurons expressing EGFP. (B) Representative image of a coronal section of a brain after double electroporation in the cortical hem and the lateral cortex. Cells targeted in the cortical hem (red) migrate tangentially, populating the marginal zone of the neocortex. Labeled cells in the ventricular zone of the cortex generate cortical projection neurons (green) that migrate radially to enter the nascent cortical plate. Scale bar = 200 µm. (C) Magnification of the area boxed in panel B displaying the lateral cortex and the two types of cells labeled after the double electroporation protocol. Dashed lines frame the marginal zone and cortical plate. Scale bar = 100 µm. High magnification on the right shows a detail of the marginal zone containing the arborized leading processes of the projection neurons intermingled with CR-cells bodies and processes. Scale bar = 10 µm. Ctx = cortex; Hem = cortical hem; MZ = marginal zone; CP = cortical plate; IZ = intermediate zone. Please click here to view a larger version of this figure.

Figure 2: Strategy B. Double electroporation strategy to study long-range interactions among cells with different spatial and temporal origin. (A) Scheme displaying the strategy to target different populations of cortical projection neurons in the lateral neocortex including the somatosensory area in different hemispheres by double in utero electroporation. Projection neurons in the somatosensory cortex of the right hemisphere targeted at E13.5 expressed BFP. Targeted projection neurons in the contralateral side targeted at E15.5 expressed nuclear EGFP (nEGFP) and membrane-targeted tdTomato (mtdTomato). (B) Representative image of a coronal section of a brain that underwent double electroporation surgery with the mentioned plasmids in panel A. Note the different distributions of the cells labeled by BFP or nEGFP and the intense labeling of the axons of upper layer callosal projection neurons (mtdTomato) labeled at E15.5. Scale bar = 500 µm. (C) Image of the somatosensory cortex located in the right hemisphere in a double electroporated brain. Note the broad distribution of the projection neurons (blue) across layers and the profuse arborization of the callosal axons (red) coming from projection neurons targeted in the contralateral hemisphere. Scale bar = 100 µm. (D) Image of the somatosensory cortex in the left hemisphere of a double electroporated brain. Note the discrete localization of the targeted projection neurons in the upper part of the cortical plate as shown by the expression of nEGFP (green), as well as the profuse red labeling surrounding cell bodies and all neuronal projections (red). Scale bar = 100 µm. (E) High magnification pictures of the somatosensory cortex in the right hemisphere showing details of the arborization of the callosal axons around targeted projection neurons. Scale bar = 10 µm. Please click here to view a larger version of this figure.

Table 1: Summary of the electroporation conditions used in the different experiments.

Table 2: Survival of embryos following double in utero electroporation in Strategy A.

Table 3: Survival of pups following double in utero electroporation in Strategy B compared with simple electroporation.

The study of cell-cell interactions in vivo in regions with high cellular density like the cerebral cortex is a complex task. Traditional approaches including the use of antibodies to label neurites are not suitable because of the lack of specific markers for different cell populations. The use of transgenic murine models, where a particular cell type expresses a fluorescent protein, is useful to visualize the neuronal processes, but this depends on the availability of such models. This task is even more complicated when trying to visualize possible differences in the interactions between two particular cell types upon perturbation of the genes of interest, because it involves the use of other animal models, like knockout mice. All of these issues made these studies complicated in the past due to economic and temporal costs.

The emergence of new techniques allowing somatic transgenesis in vivo, such as in utero electroporation, offers the possibility to design strategies like the one described in this protocol, which circumvent the use or generation of combined reporter and knockout animals, making this type of experiment more feasible. The results shown in this publication and previously published studies⁸ demonstrate that this protocol 1) successfully permits the targeting of cell populations with different temporal and spatial origin; 2) makes possible the visualization of cell-cell contacts with high resolution; and 3) is useful to detect differences in cell contacts after functional experiments.

Despite the wide use of in utero electroporation, this technique requires considerable training in order to safely perform the surgery, manipulate the embryos without damaging them, and assure the correct targeting of the desired region. However, after this training, survival of pregnant females is excellent (around 100% in our hands) and we have found no differences between single and double in utero electroporation. Single and double electroporated pregnant females recover very quickly from the surgery. In the vast majority of the cases, their behavior the day after single or double surgery seem normal and these pregnant females eat, drink, walk, and even climb without difficulties without apparent signs of pain and distress.

Survival of the embryos after the first and the second electroporation is also good overall, and the rate of abortion decreases as the age of the embryos increases (Table 2 and Table 3). The main difficulty is successful targeting in both electroporations. With older embryos from E13.5 onwards the degree of success is very high. Early ages like E11.5 are more challenging because the small size of the embryos makes handling and injection more difficult, which in addition affects their survival. However, embryos surviving the first E11.5 electroporation present very good rates of survival after the second electroporation at E13.5 (Table 2). To improve proficiency with this technique, we strongly recommend practicing surgeries in pups at ~E14.5 and progressively trying surgeries at younger ages.

Another challenge is postnatal survival, because mothers undergoing surgery do not always take care of all of their pups, although pregnant females single or double electroporated deliver the pups without problems (Table 3) and their behavior and fitness status look normal. In our hands, and with the timing described here, we find no important differences in postnatal survival after simple and double electroporation (Table 3), but to circumvent possible survival problems pups can be transferred to foster mothers upon delivery when real mothers display poor maternal behavior at birth. Providing extra nesting material and rich food to pregnant females previous to the delivery date can also help to increase the survival rates of the pups.

One of the main advantages of this strategy is the use of wild type mice for functional studies, but it can also be applied, for example, to CRE reporter mice or floxed mice for genes of interest, when available. In these mice, electroporation of CRE-recombinase expressing plasmids will allow the permanent expression of the reporter gene or the inactivation of the desired gene, respectively, in the targeted cells. For functional experiments we can also control the expression of the construct only in the cell type of interest. For example, a neuronal promoter can be used to manipulate a candidate gene only in neurons and not in neural stem cells, hence preventing undesired effects at the progenitor level. All of these considerations, together with the control of the time and the targeted region, make double in utero electroporation a very versatile technique to study cell-cell interactions not only inside the cortex but also in other structures that can be targeted using this technology.