The authors would like to thank Dr. Kathleen Mathers, Dr. Jean-Philippe Mocho and Dr. Yolanda Saavedra Torres for their help to perform in utero electroporation under aseptic procedures, and Hayley Wood for her help to prepare the drawings.
E.P. was supported by a long-term Federation of European Biochemical Societies (FEBS) fellowship and a Medical Research Council (MRC) career development fellowship, M.A.H. by a Wellcome Trust grant to Elizabeth Fisher and Victor Tybulewicz (080174/B/06/Z), H.W. by an EMBO long-term fellowship and R.A. by an MRC studentship. This work was supported by a project grant from the Wellcome Trust (086947/Z/08/Z) and by a Grant-in-Aid from the Medical Research Council (U117570528) to F.G.

In the United Kingdom, mice are housed, bred, and treated according to the guidelines approved by the Home Office under the Animal (Scientific Procedures) Act 1986.
1. Preparation: DNA Solution and Needles
<ol>
	<li>Purify plasmid DNA with an Endotoxin free Maxi-prep kit. Prepare plasmid DNA solution for injection to desired concentrations in water and add Fast Green (final concentration of 0.05%) to visualize injections. The efficiency of electroporation is highly dependent on the DNA concentration. A concentration of 1 &mu;g/&mu;l is generally used. This concentration is sufficient to visualize electroporated neurons without affecting their development. However, according to the promoter used in the plasmid vector (low expression level with cytomegalovirus promoter/enhancer, strong expression level with the cytomegalovirus immediate early enhancer and chicken &beta;-actin promoter fusion (CAG) promoter) as well as the size and the stability of the protein expressed, this concentration can be adjusted (0.25 &mu;g/&mu;l to 5 &mu;g/&mu;l).</li>
	<li>Pull glass needles using a micropipette puller.</li>
</ol>
2. Preparation of the Surgery
<ol>
	<li>Autoclave surgical instruments and phosphate buffer saline (PBS).</li>
	<li>Prepare analgesic solution in PBS (Buprenorphine, Vetergesic, final concentration of 30 &mu;g/ml). Weigh the pregnant mouse and inject subcutaneously 0.1 mg/kg of Vetergesic at least 30 minutes before the surgery. During this time, prepare the surgical area. Turn on the heating pads and the recovery chamber. Place sterile PBS in warm water bath and all sterile instruments and materials on sterile drapes. Place the platinum electrodes into a PBS filled beaker and connect to the electroporator. Fill the needle with the DNA solution using a microloader tip, connect the needle to the capillary holder and pinch off the tip of the needle with a forceps.</li>
</ol>
3. Surgery Before DNA Injection and Electroporation
<ol>
	<li>Anaesthetize a pregnant mouse (E14.5-E15.5) with isoflurane in oxygen carrier (oxygen 2 l/min) using an anaesthetic induction chamber. Wait until the animal loses righting reflex.</li>
	<li>Transfer the animal to a "pre-surgery" mask. Place a drop of eye gel on each eye to prevent corneal ulceration of the eyes while the mother is under general anesthesia. Use an electric razor to shave the hair of the abdomen. Clean the shaved area once with clorhexidine to collect flying hair.</li>
	<li>Transfer the animal to a second mask in the surgical area. Place the mouse with its back on the heating pad. Start the surgery when the pedal reflex has been lost.</li>
	<li>Put on mask and sterile gloves. Cover the animal with a sterile drape (with a small hole over the abdomen) to prevent the tissues and instruments from being contaminated by the areas of skin that have not been shaved and disinfected. Clean the shaved area at least 3 times with clorhexidine. Use a different sterile cotton swab each time. Use a scalpel to make a vertical incision along the midline (~1 inch long) through the skin. Using scissors, make a similar incision of the muscle of the abdomen along the linea alba (white line composed mostly of collagen connective tissue).</li>
	<li>Choose the most accessible embryos and place the ring forceps between two embryos and carefully pull the embryonic chain out of the abdominal cavity. From this point on, keep the embryos hydrated with sterile pre-warmed PBS.</li>
</ol>
No microscope is required for visualization.
4. Injection of DNA and Electroporation
<ol>
	<li>Start with one of the most lateral embryos, making it easier to keep track of which embryos were electroporated. Do not pull too much on the embryos as this will increase risk of hemorrhage. Manipulate the position of the embryo inside the amniotic sac using the ring forceps and stabilize the head of the embryos between the rings. Squeeze gently to push up the embryo closer to the uterine wall.</li>
	<li>With the other hand, take the capillary holder and insert the needle carefully into the middle of the hemisphere to target the lateral ventricle. Press the pedal to inject approximately 1 &mu;l of DNA solution mixed with Fast Green (less than 1 &mu;l to study dendrites). You should observe the green dye filling the lateral ventricle. Critical steps: - The sharpness of the needle is critical to pierce properly the uterine wall. It is important to minimize the movement of the needle at the surface of the uterine wall and after insertion into the ventricle because an enlargement of the hole will result 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. 
	- It is important not to inject a too large volume of DNA into the lateral ventricle because it will induce hydrocephalus (do not exceed 2 &mu;l at E14.5). The volume of DNA is adjusted according to the purpose of the experiment. If 1&mu;l is generally used for most experiments, a smaller volume of DNA (approximately 0.5 &mu;l) is required for dendrite and spine analysis. Indeed a few isolated cells need to be targeted in order to visualize and measure the dendritic arbor of electroporated neurons.</li>
	<li>Place the electrodes on the sides of the embryo head with the positive (+) paddle on the same side as the injected ventricle for cortex electroporation or on the opposite side of the injected ventricle for hippocampus electroporation (Figure 2). Then apply five 30V electrical pulses (50 msec duration) at 1 sec intervals. Critical step: Avoid applying current across the placenta as this will result in embryo death. 
	All the embryos in a pregnant mouse can be electroporated, usually with the same DNA construct to avoid any confusion. However, a long surgery decreases the survival rate of embryos. The abdominal cavity should not be opened longer than 30 min.</li>
</ol>
5. Surgery Post-electroporation
<ol>
	<li>After electroporating the embryos, add PBS into the abdominal cavity and use the ring forceps to replace the uterine horn in its original location. Suture the abdomen wall and skin with Vicryl absorbable sutures.</li>
	<li>Place the animal in a recovery chamber until it wakes up (usually 5-10 min) and then transfer in a cage placed on a heating pad.</li>
</ol>
6. Post-surgery
Check the behavior of the mice to assess pain, suffering or distress and weigh the animals 24 h and 48 h after the surgery. If needed, analgesics can be administered to minimize pain and discomfort.
7. Tissue Processing
Collect the electroporated embryos or pups at the embryonic or postnatal stages required for the experiment.
- For analysis at embryonic stages (for example to study cell proliferation or migration): 
Euthanize mother via cervical dislocation and collect the embryos. After decapitation, select the brains that have been properly electroporated, as indicated by the amount and location of the fluorescent signal, visualized across the skull using a fluorescent binocular. Dissect the brain out of the skull and fix overnight in 4% PFA and then place in 20% sucrose / PBS overnight. Embed in OCT compound, freeze at -80 &deg;C and section using a cryostat.
- For analysis at postnatal stages (for example to study dendrites and spines): 
Anesthetize pups or adult mice with intraperitoneal injection of pentobarbitone (40-60 mg/kg) and perform transcardial perfusion with PBS, followed by 4% PFA in PBS. Dissect the brains out of the skull and post-fix in 4% PFA overnight. After washings in PBS, section the brains using a vibratome (100 &mu;m sections for dendrite analysis). Mount the sections in Aqua Poly/mount using 0.16-0.19 mm thick coverslips to image dendrites and spines.
8. Representative results
Figure 3 shows examples of electroporated cells in the cerebral cortex (Figures 3A, B), in the CA1 (Figures 3C, D) and in the dentate gyrus of the hippocampus (Figures 3E, F). Wild-type mice were electroporated at E14.5 with a GFP construct (pCA-b-EGFPm5 silencer 3) and brains were harvested at postnatal day (P) 14. By injecting a small volume of DNA solution (0.5 &mu;l or less of a solution at 1 &mu;g/&mu;l), a few cells are labeled, which allows the visualization of the dendritic arborization of isolated GFP+ cells (Figure 3) as well as their spines at higher magnification (Figure 4).
<img alt="Figure 1" src="/files/ftp_upload/4163/4163fig1.jpg" /> 
Figure 1. Schematic representation of electroporation protocols that can be used to study dendrites and spines. (A) Electroporation of a GFP construct to compare dendrites and spines in wild-type and mutant mice. (B) Electroporation of GFP-shRNA (GFP is expressed from the same construct) to compare dendrites and spines in mice electroporated with a shRNA construct specific for a gene of interest (X) or a control shRNA. (C) IUE can be used together with a Cre inducible system in order to restrict expression of the shRNA to the desirable time period. In this experiment, a vector expressing a form of the Cre recombinase which can be activated by 4-hydroxytamoxifen (CAG-ERT2CreERT2; 1 &mu;g/&mu;l) 10, is electroporated together with a vector expressing a specific shRNA in a Cre dependent manner (1 &mu;g/&mu;l, and with a recombination indicator (CALNL-GFP construct, GFP expression inducible by Cre; 1 &mu;g/&mu;l) 10. The efficiency of the knockdown can be improved by increasing the concentration of the shRNA as well as increasing CAG-ERT2CreERT2 concentration.
<img alt="Figure 2" src="/files/ftp_upload/4163/4163fig2.jpg" /> 
Figure 2. Spatial control of electroporation. This figure shows where to position the paddle electrodes according to the DNA injection site in order to target the cerebral cortex or the hippocampus.
<img alt="Figure 3" src="/files/ftp_upload/4163/4163fig3.jpg" /> 
Figure 3. Visualization of the dendritic arbor of in utero electroporated cells in the cerebral cortex and hippocampus. (A, B) Coronal sections showing GFP+ pyramidal cells in the cerebral cortex, (C-D) pyramidal cells in CA1 of the hippocampus and (E, F), granule cells in the dentate gyrus at P14. A GFP construct (pCA-b-EGFPm5 silencer 3) was electroporated at E14.5. Higher magnification pictures (B, D, F) show that IUE is an efficient method to visualize dendrites. Scale bars represent 50 &mu;m (B, D and F), 150 &mu;m (A, C, E).
<img alt="Figure 4" src="/files/ftp_upload/4163/4163fig4.jpg" /> 
Figure 4. Visualization of dendritic spines in P14 neurons that were electroporated in utero at E14.5 with a GFP expressing construct. (A, B) High magnification images of spines from basal dendrites of hippocampal pyramidal neurons. Scale bars represent 5 &mu;m (A) and 2 &mu;m (B).

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Biol. 240, 237-246 (2001).</li><li>Shimogori, T., Ogawa, M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Gene+application+with+in+utero+electroporation+in+mouse+embryonic+brain.">Gene application with in utero electroporation in mouse embryonic brain.</a> Dev. 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No conflicts of interest declared.

IUE is a powerful tool to manipulate gene expression not only in space but also in time. We show here that this technique can be used to visualize and genetically manipulate dendrites and spines in the cerebral cortex and hippocampus of mice. Besides the advantages previously cited, it is worth noting that IUE, in contrast to Golgi method, can be combined with immunohistochemistry or in situ hybridization, which allows for example to phenotype the electroporated cells. It is also important to mention that this procedure does not induce evident brain malformations despite its relative invasiveness. In addition, at the cellular level, IUE does not modify the electrophysiological properties of the electroporated neurons13. While our demonstration focuses on the visualization of dendrite and spine morphologies, IUE of cortical or hippocampal neurons at E14.5 could also be used to study other developmental events such as axon formation and guidance. In addition, the same kind of protocol could be implemented at other stages of embryonic development to target different populations. For example, a developmentally very late cortical electroporation paradigm at E18.5 can be performed to drive expression in astrocytic progenitors 1. Similarly, while an electroporation of the hippocampus at E14.5 allows to target CA1-CA3 pyramidal neuron progenitors and dentate granule cell progenitor at the same time, a late hippocampal electroporation (E18.5 or early postnatal) would allow to target different dentate granule progenitors 14. In this case, the injected volume of DNA can be increased as well as the intensity of current. 
Transgenes introduced by IUE appear to remain episomal and are therefore lost from cells following successive cell divisions. In postmitotic cells such as neurons, however, the episomal transgenes remain active for months after electroporation allowing long-term studies 13,15. In our study, we have observed bright GFP+ cells up to 7 weeks after birth (the latest time point we analyzed) indicating that embryonic targeting of cortical or hippocampal neuronal precursors using IUE results in persistent expression of the transgene from early developmental time points up to adulthood.
A current limitation of the technique is that it is difficult to exert a fine control over the total number of electroporated cells. However, by decreasing the injected volume of DNA solution, we have shown that it is possible to label a few cells and to visualize the dendritic arborization of isolated GFP+ cells as well as their spines. The dimension of the transfected area could also be adjusted by modifying the parameters of the electroporation such as intensity of current and number of pulses or the diameter of the electroporation paddles.
Altogether IUE is a method that is easy to implement, rapid and efficient to study dendrites and spines in vivo.

<table border="1">
 <tbody>
 <tr>
 <td>Name of the reagent</td>
 <td>Company</td>
 <td>Catalogue number</td>
 <td>Comments </td>
 </tr>
 <tr>
 <td>&nbsp;</td>
 <td>&nbsp;</td>
 <td>&nbsp;</td>
 <td colspan="4">Preparation of needles and DNA solution for injection</td>
 </tr>
 <tr>
 <td>Endofree Plasmid Maxi Kit</td>
 <td>Qiagen</td>
 <td>12362</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>Fast Green</td>
 <td>Sigma</td>
 <td>F-7258</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>Borosilicate glass capillaries 
 1.0 mm O.D. x 0.58 mm I.D.</td>
 <td>Harvard Apparatus</td>
 <td>30-0016</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>Microloader tips</td>
 <td>Eppendorf</td>
 <td>5242956003</td>
 <td>For Eppendorf pipettes 0.5 &mu;l-10 &mu;l / 2-20 &mu;l</td>
 </tr>
 <tr>
 <td>&nbsp;</td>
 <td>&nbsp;</td>
 <td>&nbsp;</td>
 <td>Material for surgery</td>
 </tr>
 <tr>
 <td>Extra thin Iris scissors </td>
 <td>Fine Science Tools</td>
 <td>14088-10</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>Curved Forceps</td>
 <td>Fine Science Tools</td>
 <td>91197-00</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>Ring forceps</td>
 <td>Fine Science Tools</td>
 <td>11103-09</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>Needle holder</td>
 <td>Fine Science Tools</td>
 <td>12002-12</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>Graefe Forceps </td>
 <td>Fine Science Tools</td>
 <td>11050-10</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>Vicryl absorbable suture</td>
 <td>Ethicon Inc (Johnson &amp; Johnson)</td>
 <td>W9074</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>Sterile drapes 30cm x 45 cm</td>
 <td>Buster</td>
 <td>141765</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>Sterile swabs</td>
 <td>Shermond</td>
 <td>HUBY-340</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>Cotton buds</td>
 <td>Clean Cross Co., Ltd</td>
 <td>1860</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>Buprenorphine (Vetergesic)</td>
 <td>Alstoe Animal Health</td>
 <td>&nbsp;</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>Clorhexidine </td>
 <td>Vetasept</td>
 <td>XHG007</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>Eye gel Viscotears </td>
 <td>Novartis </td>
 <td>&nbsp;</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>Isoflurane</td>
 <td>Abbott Laboratories</td>
 <td>B506</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>Pentoject, Pentobarbitone Sodium 20%</td>
 <td>Animalcare</td>
 <td>&nbsp;</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>&nbsp;</td>
 <td>&nbsp;</td>
 <td>&nbsp;</td>
 <td colspan="4">Electroporation</td>
 </tr>
 <tr>
 <td>Electroporator </td>
 <td>BTX</td>
 <td>ECM830 </td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>Platinum Tweezertrode 5mm</td>
 <td>BTX, Harvard Apparatus</td>
 <td>45-0489</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>Femtojet microinjector</td>
 <td>Eppendorf</td>
 <td>5247000030</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>Foot Control for Femtojet Microinjector</td>
 <td>Eppendorf</td>
 <td>5247623002</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>Capillary holder</td>
 <td>Eppendorf</td>
 <td>5176190002</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>&nbsp;</td>
 <td>&nbsp;</td>
 <td>&nbsp;</td>
 <td>Tissue processing</td>
 </tr>
 <tr>
 <td>Paraformaldehyde</td>
 <td>Sigma</td>
 <td>P6148</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>Sucrose</td>
 <td>VWR (Prolabo)</td>
 <td>27480.294</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>Microscope slides</td>
 <td>ThermoScientific (Menzel-Gl&auml;ser)</td>
 <td>J1800AMNZ</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>Coverslips</td>
 <td>Menzel-Gl&auml;ser </td>
 <td>&nbsp;</td>
 <td>22 x 50 mm #1,5</td>
 </tr>
 <tr>
 <td>Aqua Poly/mount</td>
 <td>Polysciences, Inc</td>
 <td>18606</td>
 <td>&nbsp;</td>
 </tr>
 </tbody>
</table>

visualization and genetic manipulation of dendrites and spines in the mouse cerebral cortex and hippocampus using in utero electroporation

In utero electroporation (IUE) has become a powerful technique to study the development of different regions of the embryonic nervous system 1-5. To date this tool has been widely used to study the regulation of cellular proliferation, differentiation and neuronal migration especially in the developing cerebral cortex 6-8. Here we detail our protocol to electroporate in utero the cerebral cortex and the hippocampus and provide evidence that this approach can be used to study dendrites and spines in these two cerebral regions.
Visualization and manipulation of neurons in primary cultures have contributed to a better understanding of the processes involved in dendrite, spine and synapse development. However neurons growing in vitro are not exposed to all the physiological cues that can affect dendrite and/or spine formation and maintenance during normal development. Our knowledge of dendrite and spine structures in vivo in wild-type or mutant mice comes mostly from observations using the Golgi-Cox method 9. However, Golgi staining is considered to be unpredictable. Indeed, groups of nerve cells and fiber tracts are labeled randomly, with particular areas often appearing completely stained while adjacent areas are devoid of staining. Recent studies have shown that IUE of fluorescent constructs represents an attractive alternative method to study dendrites, spines as well as synapses in mutant / wild-type mice 10-11 (Figure 1A). Moreover in comparison to the generation of mouse knockouts, IUE represents a rapid approach to perform gain and loss of function studies in specific population of cells during a specific time window. In addition, IUE has been successfully used with inducible gene expression or inducible RNAi approaches to refine the temporal control over the expression of a gene or shRNA 12. These advantages of IUE have thus opened new dimensions to study the effect of gene expression/suppression on dendrites and spines not only in specific cerebral structures (Figure 1B) but also at a specific time point of development (Figure 1C).
Finally, IUE provides a useful tool to identify functional interactions between genes involved in dendrite, spine and/or synapse development. Indeed, in contrast to other gene transfer methods such as virus, it is straightforward to combine multiple RNAi or transgenes in the same population of cells. 
In summary, IUE is a powerful method that has already contributed to the characterization of molecular mechanisms underlying brain function and disease and it should also be useful in the study of dendrites and spines.

Watch this Scientific Journal Video about Visualization and Genetic Manipulation of Dendrites and Spines in the Mouse Cerebral Cortex and Hippocampus using In utero Electroporation at JoVE.com

Visualization and Genetic Manipulation of Dendrites and Spines in the Mouse Cerebral Cortex and Hippocampus using In utero Electroporation

This article describes in detail a protocol to electroporate in utero the cerebral cortex and the hippocampus at E14.5 in mice. We also show that this is a valuable method to study dendrites and spines in these two cerebral regions.

Visualization and Genetic Manipulation of Dendrites and Spines in the Mouse Cerebral Cortex and Hippocampus using In utero Electroporation

In utero electroporation (IUE) has become a powerful technique to study the development of different regions of the embryonic nervous system 1-5. To date ...

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JoVE Journal

Neuroscience

17.8K Views.  MRC National Institute for Medical Research. This article describes in detail a protocol to electroporate in utero the cerebral cortex and the hippocampus at E14.5 in mice. We also show that this is a valuable method to study dendrites and spines in these two cerebral regions.

Video: Visualization and Genetic Manipulation of Dendrites and Spines in the Mouse Cerebral Cortex and Hippocampus using In utero Electroporation

Lentivirus-mediated Genetic Manipulation and Visualization of Olfactory Sensory Neurons in vivo

Development of a precise olfactory circuit relies on accurate projection of olfactory sensory neuron (OSN) axons to their synaptic 
targets in the olfactory bulb (OB). The molecular mechanisms of OSN axon growth and targeting are not well understood. Manipulating gene expression and subsequent 
visualizing of single OSN axons and their terminal arbor morphology have thus far been challenging. To study gene function at the single cell level within a specified 
time frame, we developed a lentiviral based technique to manipulate gene expression in OSNs in vivo. Lentiviral particles are delivered to OSNs by microinjection 
into the olfactory epithelium (OE). Expression cassettes are then permanently integrated into the genome of transduced OSNs. Green fluorescent protein expression 
identifies infected OSNs and outlines their entire morphology, including the axon terminal arbor. Due to the short turnaround time between microinjection and 
reporter detection, gene function studies can be focused within a very narrow period of development. With this method, we have detected GFP expression within as 
few as three days and as long as three months following injection. We have achieved both over-expression and shRNA mediated knock-down by lentiviral microinjection. 
This method provides detailed morphologies of OSN cell bodies and axons at the single cell level in vivo, and thus allows characterization of candidate gene function 
during olfactory development.

Lentivirus-mediated Genetic Manipulation and Visualization of Olfactory Sensory Neurons in vivo

We present a lentiviral technique for genetic manipulation and visualization of single olfactory sensory neuron axon and its terminal arborization in vivo.

Development of a precise olfactory circuit relies on accurate projection of olfactory sensory neuron (OSN) axons to their synaptic 
targets in the ...

Visualization of Proprioceptors in Drosophila Larvae and Pupae

Proprioception is the ability to sense the motion, or position, of body parts by responding to stimuli arising within the body. In fruitflies and other insects proprioception is provided by specialized sensory organs termed chordotonal organs (ChOs) 2. Like many other organs in Drosophila, ChOs develop twice during the life cycle of the fly. First, the larval ChOs develop during embryogenesis. Then, the adult ChOs start to develop in the larval imaginal discs and continue to differentiate during metamorphosis.
The development of larval ChOs during embryogenesis has been studied extensively 10,11,13,15,16. The centerpiece of each ChO is a sensory unit composed of a neuron and a scolopale cell. The sensory unit is stretched between two types of accessory cells that attach to the cuticle via specialized epidermal attachment cells 1,9,14. When a fly larva moves, the relative displacement of the epidermal attachment cells leads to stretching of the sensory unit and consequent opening of specific transient receptor potential vanilloid (TRPV) channels at the outer segment of the dendrite 8,12. The elicited signal is then transferred to the locomotor central pattern generator circuit in the central nervous system.
Multiple ChOs have been described in the adult fly 7. These are located near the joints of the adult fly appendages (legs, wings and halters) and in the thorax and abdomen. In addition, several hundreds of ChOs collectively form the Johnston's organ in the adult antenna that transduce acoustic to mechanical energy 3,5,17,4.
In contrast to the extensive knowledge about the development of ChOs in embryonic stages, very little is known about the morphology of these organs during larval stages. Moreover, with the exception of femoral ChOs 18 and Johnston's organ, our knowledge about the development and structure of ChOs in the adult fly is very fragmentary.
Here we describe a method for staining and visualizing ChOs in third instar larvae and pupae. This method can be applied together with genetic tools to better characterize the morphology and understand the development of the various ChOs in the fly.

Visualization of Proprioceptors in Drosophila Larvae and Pupae

A method to immunostain and visualize chordotonal organs in larvae and pupae of Drosophila melanogaster is described.

Proprioception is the ability to sense the motion, or position, of body parts by responding to stimuli arising within the body. In fruitflies and other ...

Voltage-sensitive Dye Recording from Axons, Dendrites and Dendritic Spines of Individual Neurons in Brain Slices

Understanding the biophysical properties and functional organization of single neurons and how they process information is fundamental for understanding how the brain works. The primary function of any nerve cell is to process electrical signals, usually from multiple sources. Electrical properties of neuronal processes are extraordinarily complex, dynamic, and, in the general case, impossible to predict in the absence of detailed measurements. To obtain such a measurement one would, ideally, like to be able to monitor, at multiple sites, subthreshold events as they travel from the sites of origin on neuronal processes and summate at particular locations to influence action potential initiation. This goal has not been achieved in any neuron due to technical limitations of measurements that employ electrodes. To overcome this drawback, it is highly desirable to complement the patch-electrode approach with imaging techniques that permit extensive parallel recordings from all parts of a neuron. Here, we describe such a technique - optical recording of membrane potential transients with organic voltage-sensitive dyes (Vm-imaging) - characterized by sub-millisecond and sub-micrometer resolution. Our method is based on pioneering work on voltage-sensitive molecular probes 2. Many aspects of the initial technology have been continuously improved over several decades 3, 5, 11. Additionally, previous work documented two essential characteristics of Vm-imaging. Firstly, fluorescence signals are linearly proportional to membrane potential over the entire physiological range (-100 mV to +100 mV; 10, 14, 16). Secondly, loading neurons with the voltage-sensitive dye used here (JPW 3028) does not have detectable pharmacological effects. The recorded broadening of the spike during dye loading is completely reversible 4, 7. Additionally, experimental evidence shows that it is possible to obtain a significant number (up to hundreds) of recordings prior to any detectable phototoxic effects 4, 6, 12, 13. At present, we take advantage of the superb brightness and stability of a laser light source at near-optimal wavelength to maximize the sensitivity of the Vm-imaging technique. The current sensitivity permits multiple site optical recordings of Vm transients from all parts of a neuron, including axons and axon collaterals, terminal dendritic branches, and individual dendritic spines. The acquired information on signal interactions can be analyzed quantitatively as well as directly visualized in the form of a movie.

An imaging technique for monitoring of membrane potential changes with sub-micrometer spatial and sub-millisecond temporal resolution is described. The technique, based on laser excitation of voltage-sensitive dyes, allows measurements of signals in axons and axon collaterals, terminal dendritic branches, and individual dendritic spines.

Understanding the biophysical properties and functional organization of single neurons and how they process information is fundamental for understanding ...

Simultaneous Electroencephalography, Real-time Measurement of Lactate Concentration and Optogenetic Manipulation of Neuronal Activity in the Rodent Cerebral Cortex

Although the brain represents less than 5% of the body by mass, it utilizes approximately one quarter of the glucose used by the body at rest1. The function of non rapid eye movement sleep (NREMS), the largest portion of sleep by time, is uncertain. However, one salient feature of NREMS is a significant reduction in the rate of cerebral glucose utilization relative to wakefulness2-4. This and other findings have led to the widely held belief that sleep serves a function related to cerebral metabolism. Yet, the mechanisms underlying the reduction in cerebral glucose metabolism during NREMS remain to be elucidated.
	One phenomenon associated with NREMS that might impact cerebral metabolic rate is the occurrence of slow waves, oscillations at frequencies less than 4 Hz, in the electroencephalogram5,6. These slow waves detected at the level of the skull or cerebral cortical surface reflect the oscillations of underlying neurons between a depolarized/up state and a hyperpolarized/down state7. During the down state, cells do not undergo action potentials for intervals of up to several hundred milliseconds. Restoration of ionic concentration gradients subsequent to action potentials represents a significant metabolic load on the cell8; absence of action potentials during down states associated with NREMS may contribute to reduced metabolism relative to wake.
	Two technical challenges had to be addressed in order for this hypothetical relationship to be tested. First, it was necessary to measure cerebral glycolytic metabolism with a temporal resolution reflective of the dynamics of the cerebral EEG (that is, over seconds rather than minutes). To do so, we measured the concentration of lactate, the product of aerobic glycolysis, and therefore a readout of the rate of glucose metabolism in the brains of mice. Lactate was measured using a lactate oxidase based real time sensor embedded in the frontal cortex. The sensing mechanism consists of a platinum-iridium electrode surrounded by a layer of lactate oxidase molecules. Metabolism of lactate by lactate oxidase produces hydrogen peroxide, which produces a current in the platinum-iridium electrode. So a ramping up of cerebral glycolysis provides an increase in the concentration of substrate for lactate oxidase, which then is reflected in increased current at the sensing electrode. It was additionally necessary to measure these variables while manipulating the excitability of the cerebral cortex, in order to isolate this variable from other facets of NREMS.
	We devised an experimental system for simultaneous measurement of neuronal activity via the elecetroencephalogram, measurement of glycolytic flux via a lactate biosensor, and manipulation of cerebral cortical neuronal activity via optogenetic activation of pyramidal neurons. We have utilized this system to document the relationship between sleep-related electroencephalographic waveforms and the moment-to-moment dynamics of lactate concentration in the cerebral cortex. The protocol may be useful for any individual interested in studying, in freely behaving rodents, the relationship between neuronal activity measured at the electroencephalographic level and cellular energetics within the brain.

A procedure is described for manipulating the activity of cerebral cortical pyramidal neurons optogenetically while the electroencephalogram, electromyogram, and cerebral lactate concentration are monitored. Experimental recordings are performed on cable-tethered mice while they undergo spontaneous sleep/wake cycles. Optogenetic equipment is assembled in our laboratory; recording equipment is commercially available.

Although the brain represents less than 5% of the body  by mass, it utilizes approximately one quarter of the glucose used by the body  at rest1. The ...

Dual Electrophysiological Recordings of Synaptically-evoked Astroglial and Neuronal Responses in Acute Hippocampal Slices

Astrocytes form together with neurons tripartite synapses, where they integrate and modulate neuronal activity. Indeed, astrocytes sense neuronal inputs through activation of their ion channels and neurotransmitter receptors, and process information in part through activity-dependent release of gliotransmitters. Furthermore, astrocytes constitute the main uptake system for glutamate, contribute to potassium spatial buffering, as well as to GABA clearance. These cells therefore constantly monitor synaptic activity, and are thereby sensitive indicators for alterations in synaptically-released glutamate, GABA and extracellular potassium levels. Additionally, alterations in astroglial uptake activity or buffering capacity can have severe effects on neuronal functions, and might be overlooked when characterizing physiopathological situations or knockout mice. Dual recording of neuronal and astroglial activities is therefore an important method to study alterations in synaptic strength associated to concomitant changes in astroglial uptake and buffering capacities. Here we describe how to prepare hippocampal slices, how to identify stratum radiatum astrocytes, and how to record simultaneously neuronal and astroglial electrophysiological responses. Furthermore, we describe how to isolate pharmacologically the synaptically-evoked astroglial currents.

The preparation of acute brain slices from isolated hippocampi, as well as the simultaneous electrophysiological recordings of astrocytes and neurons in stratum radiatum during stimulation of schaffer collaterals is described. The pharmacological isolation of astroglial potassium and glutamate transporter currents is demonstrated.

Astrocytes form together with neurons tripartite synapses, where they integrate and modulate neuronal activity. Indeed, astrocytes sense neuronal inputs ...

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 ...

Two-Photon in vivo Imaging of Dendritic Spines in the Mouse Cortex Using a Thinned-skull Preparation

In the mammalian cortex, neurons form extremely complicated networks and exchange information at synapses. Changes in synaptic strength, as well as addition/removal of synapses, occur in an experience-dependent manner, providing the structural foundation of neuronal plasticity. As postsynaptic components of the most excitatory synapses in the cortex, dendritic spines are considered to be a good proxy of synapses. Taking advantages of mouse genetics and fluorescent labeling techniques, individual neurons and their synaptic structures can be labeled in the intact brain. Here we introduce a transcranial imaging protocol using two-photon laser scanning microscopy to follow fluorescently labeled postsynaptic dendritic spines over time in vivo. This protocol utilizes a thinned-skull preparation, which keeps the skull intact and avoids inflammatory effects caused by exposure of the meninges and the cortex. Therefore, images can be acquired immediately after surgery is performed. The experimental procedure can be performed repetitively over various time intervals ranging from hours to years. The application of this preparation can also be expanded to investigate different cortical regions and layers, as well as other cell types, under physiological and pathological conditions.

Two-Photon in vivo Imaging of Dendritic Spines in the Mouse Cortex Using a Thinned-skull Preparation

Time-lapse imaging in the living animal provides valuable information on structural reorganization in the intact brain. Here, we introduce a thinned-skull preparation that allows transcranial imaging of fluorescently labeled synaptic structures in the living mouse cortex by two-photon microscopy.

In the mammalian cortex, neurons form extremely complicated networks and exchange information at synapses. Changes in synaptic strength, as well as ...

Cortex-, Hippocampus-, Thalamus-, Hypothalamus-, Lateral Septal Nucleus- and Striatum-specific In Utero Electroporation in the C57BL/6 Mouse

In utero electroporation is a widely used technique for fast and efficient spatiotemporal manipulation of various genes in the rodent central nervous system. Overexpression of desired genes is just as possible as shRNA mediated loss-of-function studies. Therefore it offers a wide range of applications. The feasibility to target particular cells in a distinct area further increases the range of potential applications of this very useful method. For efficiently targeting specific regions knowledge about the subtleties, such as the embryonic stage, the voltage to apply and most importantly the position of the electrodes, is indispensable.
Here, we provide a detailed protocol that allows for specific and efficient in utero electroporation of several regions of the C57BL/6 mouse central nervous system. In particular it is shown how to transfect regions the develop into the retrosplenial cortex, the motor cortex, the somatosensory cortex, the piriform cortex, the cornu ammonis 1-3, the dentate gyrus, the striatum, the lateral septal nucleus, the thalamus and the hypothalamus. For this information about the appropriate embryonic stage, the appropriate voltage for the corresponding embryonic stage is provided. Most importantly an angle-map, which indicates the appropriate position of the positive pole, is depicted. This standardized protocol helps to facilitate efficient in utero electroporation, which might also lead to a reduced number of animals.

Cortex-, Hippocampus-, Thalamus-, Hypothalamus-, Lateral Septal Nucleus- and Striatum-specific In Utero Electroporation in the C57BL/6 Mouse

This protocol describes in detail how to specifically transfect different regions in the C57BL/6 central nervous system via in utero electroporation. Included in this protocol are detailed instructions for transfections of regions that develop into the cortex, hippocampus, thalamus, hypothalamus, lateral septal nucleus and striatum.

In utero electroporation is a widely used technique for fast and efficient spatiotemporal manipulation of various genes in the rodent central nervous ...

Inducing Cre-lox Recombination in Mouse Cerebral Cortex Through In Utero Electroporation

Cell-autonomous neuronal functions of genes can be revealed by causing loss or gain of function of a gene in a small and sparse population of neurons. To do so requires generating a mosaic in which neurons with loss or gain of function of a gene are surrounded by genetically unperturbed tissue. Here, we combine the Cre-lox recombination system with in utero electroporation in order to generate mosaic brain tissue that can be used to study the cell-autonomous function of genes in neurons. DNA constructs (available through repositories), coding for a fluorescent label and Cre recombinase, are introduced into developing cortical neurons containing genes flanked with loxP sites in the brains of mouse embryos using in utero electroporation. Additionally, we describe various adaptations to the in utero electroporation method that increase survivability and reproducibility. This method also involves establishing a titer for Cre-mediated recombination in a sparse or dense population of neurons. Histological preparations of labeled brain tissue do not require (but can be adapted to) immunohistochemistry. The constructs used guarantee that fluorescently labeled neurons carry the gene for Cre recombinase. Histological preparations allow morphological analysis of neurons through confocal imaging of dendritic and axonal arbors and dendritic spines. Because loss or gain of function is achieved in sparse mosaic tissue, this method permits the study of cell-autonomous necessity and sufficiency of gene products in vivo.

Inducing Cre-lox Recombination in Mouse Cerebral Cortex Through In Utero Electroporation

Cell-autonomous functions of genes in the brain can be studied by inducing loss or gain of function in sparse populations of cells. Here, we describe in utero electroporation to deliver Cre recombinase into sparse populations of developing cortical neurons with floxed genes to cause loss of function in vivo.

Cell-autonomous neuronal functions of genes can be revealed by causing loss or gain of function of a gene in a small and sparse population of neurons. To ...

Rapid Golgi Stain for Dendritic Spine Visualization in Hippocampus and Prefrontal Cortex

Golgi impregnation, using the Golgi staining kit with minor adaptations, is used to impregnate dendritic spines in the rat hippocampus and medial prefrontal cortex. This technique is a marked improvement over previous methods of Golgi impregnation because the premixed chemicals are safer to use, neurons are consistently well impregnated, there is far less background debris, and for a given region, there are extremely small deviations in spine density between experiments. Moreover, brains can be accumulated after a certain point and kept frozen until further processing. Using this method any brain region of interest can be studied. Once stained and cover slipped, dendritic spine density is determined by counting the number of spines for a length of dendrite and expressed as spine density per 10 &#181;m dendrite.

The protocol describes a modification of the rapid Golgi method, which can be adapted to any part of the nervous system, for staining neurons in the hippocampus and medial prefrontal cortex of the rat.