C.M.P. acknowledges funding from a Whitaker International Scholar grant administered by the Institute for International Education. A.K. was sponsored by the Marie Curie IEF (No. 625372). A.W. acknowledges funding from the European Research Council (ERC) under the European Union&#8217;s Horizon 2020 research and innovation programme (grant agreement No. 716867). A.W. additionally acknowledges the Excellence Initiative of Aix-Marseille University - A*MIDEX, a French &#8220;Investissements d&#8217;Avenir&#8221; programme. The authors acknowledge Dr. Ilke Uguz, Dr. Sahika Inal, Dr. Vincenzo Curto, Dr. Mary Donahue, Dr. Marc Ferro, and Zs&#243;fia Magl&#243;czky for their participation in fruitful discussions.

All experimental procedures were performed according to the ethical guidelines of the Institut de Neurosciences des Syst&#232;mes and approved by the local Ethical Committees and Veterinary Offices.
NOTE: Seventeen adult male OF1 mice were used for the experiments. Mice were entrained to a 12 h light/dark cycle with food and water available ad libitum.
1. Anesthesia
<ol>
	<li>Inject intraperitoneally a mixture of ketamine and xylazine (100 mg/kg body weight and 10 mg/kg body weight, respectively) to anesthetize the animal.</li>
	<li>Check the level of anesthesia by observing the respiratory rate and whisking and by checking the mouse&#8217;s response to pain. 
	NOTE: When the mouse&#8217;s breathing becomes regular, no whisking can be observed, and the animal does not react to tail pinches, the anesthesia is deep enough to continue.
	<ol>
		<li>Place the animal on an electrically programmable heating pad. Cover the rectal temperature probe with a petroleum jelly-based product (see Table of Materials) and place it gently into the rectum (1&#8211;2 cm deep) of the mouse to monitor its body temperature. Maintain a body temperature between 36.5 and 37.5 &#176;C during the surgical procedures and experimental recordings.</li>
		<li>Monitor the anesthesia level by checking the mouse&#8217;s reflexes, whisker movements, and its frequency of breathing. Taking into account the level of anesthesia recorded at least every 30 min, give a small dose of a ketamine-xylazine cocktail (20&#8211;50 &#181;L, the same concentration used as before) intramuscularly.</li>
	</ol>
	</li>
</ol>
2. Surgery/Craniotomy
<ol>
	<li>Fix the head of the mouse in a stereotaxic frame. Using a 30 G needle, inject local analgesic ropivacaine (5 &#181;L, 7.5 mg/mL, see Table of Materials) subcutaneously at the planned incision site. Allow 5 min for it to take effect.</li>
	<li>Make a straight cut midline in the skin above the skull with a scalpel. Gently pull the skin toward the sides with fine forceps and clamp it aside with bulldog serrefine clamps to leave the skull exposed for further work.</li>
	<li>Clean the skull of fascia with a scalpel or any similar tool. In case of superficial bleeding, remove the blood with cotton swabs or small pieces of paper towel.</li>
	<li>Take a poly(3,4-ethylenedioxythiophene) polystyrene sulfonate (PEDOT:PSS)-coated ground screw (size: #00, diameter: 0.047 in, length: 1/8 in, see Table of Materials) with a soldered wire and connect it with a connector to the amplifier headstage.</li>
	<li>Moisten the skull at the desired hole site and drill a hole at high speed using a fine, round drill bit (with a 0.4 mm diameter) on the skull above the cerebellum until the dura is visible. Put the ground screw into the hole and screw it in with a precision screwdriver until it reaches the top of the cerebellum. 
	NOTE: The head screw was dip-coated with a PEDOT:PSS solution containing 1% 3-glycidyloxypropyl)trimethoxysilane (GOPS) by weight followed by baking at 140 &#176;C for 90 min. PEDOT:PSS is a conjugated polymer with a volumetric capacitance that is known to be biocompatible. GOPS is a cross-linker mixed with PEDOT:PSS to increase stability in aqueous media (Figure 2).</li>
	<li>With the help of the stereotaxic frame, measure the stereotaxic coordinates for the desired brain region. For example, the region of interest is the hippocampus, anteroposterior (AP) -1.8 mm and mediolateral (ML) 1.8 mm from the Bregma point based on the brain atlas for mice32. 
	NOTE: These are coordinates for the right hemisphere (Figure 2).</li>
	<li>Thin an approximately 1 to 2 mm&#160;diameter area of the skull above the target region using a reliable dental drill (see Table of materials) set at a fast speed until a thin, well-polished, transparent bone membrane remains.</li>
	<li>Then, if the thickness of the bone membrane is thin enough (&#60;200 &#181;m), make a small hole with thin forceps and gently remove the thin layer of the bone33. Use custom-made hook-tipped needle to remove the dura. Minimize the size of craniotomy and durotomy to prevent the development of edemas and to minimize cardiac and/or respiratory pulsations of the brain. 
	NOTE: The craniotomy must be filled with a droplet of saline solution to prevent drying and then regularly refilled during the experiment (Figure 2).</li>
</ol>
3. Insertion of the Multichannel Silicon Probe
<ol>
	<li>Use stereotaxic arms in a slight AP angle (20&#176;) for the silicon probe to leave ample space for the positioning of the other two implants and to have the recording and the injection sites of the electrode, ion pump, and micropipette as close as possible. 
	NOTE: Electrodes, syringes, and ion pumps were covered with a drop of DiI stain solution (1,1&#39;-dioctadecyl-3,3,3&#39;,3&#39;-tetramethylindocarbocyanine perchlorate [DiI]), for the post hoc visualization of the implantation traces (0.5 mg/ml DiI in dimethyl sulfoxide).</li>
	<li>Place the silicon probe on the stereotaxic arm attached to a magnetic holder and place it next to the stereotaxic frame. Set the AP angle (20&#176;), and then connect the probe to the headstage and to the ground screw.</li>
	<li>Slowly lower the silicon probe into the hippocampus with the help of the micron-precise stereotaxic arm or a motorized micromanipulator to avoid lateral movements (Figure 2 and Figure 3).
	<ol>
		<li>Initiate the recording software and record&#8212;with the headstage, the connected amplifier, and a computer&#8212;electric neuronal signals while moving the multichannel silicon probe from the top of the cortex until the targeted dorsoventral (DV) position is reached (-1,800 &#181;m from the cortical surface). Record and watch the local field potential signal (LFP) during penetration on the computer screen. 
		NOTE: Control the descent of the probe so that it is moving slowly and continuously while recording, to have better visual control for the penetration and for reaching the target zone.</li>
		<li>Use the ripple activity in the pyramidal layer of the hippocampal formation in the recorded LFP as a marker of the target zone. 
		NOTE: Ripple activity is visible on one or two neighboring channels of the multichannel silicon (Si) probe having a 100 &#181;m distance between recording sites (Figure 4).</li>
		<li>Record LFP signals from the layers of the cortex and the hippocampus simultaneously through the multichannel amplifier&#8217;s software (see Table of Materials) with the help of the multichannel Si probes (Figure 4).</li>
	</ol>
	</li>
</ol>
4. Insertion of &#181;FIP
<ol>
	<li>Connect tubes (see Table of Materials) to the inlet of the &#181;FIP and fill the probe with 0.05 M GABA solution. Remove the tubes and close the inlet with paraffin film wrapping. Connect electrical leads to the source measurement unit.</li>
	<li>Insert the &#181;FIP with the help of the stereotaxic arm at a mediolateral (MP) angle (20&#176;). The Si probe remains inserted during the whole process. 
	NOTE: &#181;FIP is very flexible and may benefit from the support of a small and clean paintbrush to keep it straight until it reaches the brain surface. After that step, &#181;FIP can be lowered gently with axial movements.</li>
	<li>Lower the &#181;FIP slowly with axial movements and never let it bend during the trajectory until it reaches the dorsoventral (DV) coordinate (-1,200 &#181;m from the cortical surface). 
	NOTE: Try to put the two devices (&#181;FIP and silicon probe) as close to each other as possible, considering the 300 &#956;m distance of the outlet from the &#181;FIP tip. 
	CAUTION: Avoid any mechanical issues among the devices and their connectors during insertion (Figure 2B and Figure 3B).</li>
</ol>
5. Preparation of devices for Seizure Induction
<ol>
	<li>Change the metal needle of the syringe (10 &#181;L) (see Table of Materials). Remove the needle-holding metal part, place and fix the micropipette (outer diameter [OD]: 1.2 mm, inner diameter [ID]: 0.75 mm, tip diameter: 20&#8211;50 &#181;m with &#177; 0.5 cm of tapering of the shank), and then replace the needle-holding element.</li>
	<li>Position the syringe and the attached borosilicate micropipette at a 20&#176; lateromedial (LM) angle for the injection of 4AP (50 mM in artificial cerebrospinal fluid [ACSF]). 
	CAUTION: Do not use the metal needle of the syringe or a micropipette with a tip bigger than 50 &#181;m.</li>
	<li>Draw 500 nL&#8211;1 &#181;L of 50 mM 4AP with the help of an automated microinjection pump.</li>
</ol>
6. Insertion of the Glass Pipette Attached to a Syringe for 4AP Injection
<ol>
	<li>Lower the glass micropipette attached to the syringe to the aimed DV position (-1,500 &#181;m), and then inject 250 nL of the 4AP solution (Figure 2 and Figure 3). Start recording with the recording software. Watch the screen and wait for the first interictal spike to appear.</li>
	<li>Start the GABA delivery by &#181;FIP immediately with the appearance of the first interictal spike. Deliver GABA by applying 1 V between source and target for 100 s followed by 1 s off, for 30 cycles. With the help of the recording software, record for a minimum of 2 h. 
	NOTE: The total mass of the delivered GABA is around 1 nmol (Figure 5).</li>
	<li>At the end of the experiment, gently remove the inserted probes and the ground screw, and remove the animal from the stereotaxic equipment.&#160;Animals were euthanized using an overdose of drug (i.p.100mg/kg pentobarbital). Death was confirmed by cessation of breath and circulation.</li>
</ol>
7. Evaluation of the Placement of the Implants
<ol>
	<li>After euthanizing the animal, perfuse it transcardially, first with 50 mL of saline and then with 150 mL of an ice-cold fixative solution containing 4% paraformaldehyde (PFA) in 0.1 M phosphate buffer (PB)34. 
	CAUTION: PFA is hazardous and must be handled with care.</li>
	<li>Decapitate the animal, and then remove the skin and the muscle from the top and sides of the skull. Starting from the foramen magnum, make lateral incisions in the skull toward the ears and a sagittal midline incision, taking great care not to damage the brain. Gently remove the skull with a bone trimmer. Remove the brain, and then cut a tissue block from the region of interest (from the Bregma point, -1 to -3 mm AP) with the help of a brain matrix (see Table of Materials).</li>
	<li>Glue the tissue block to the specimen holder of a vibratome, put the stand into it, and set the vibratome to 40 &#181;m thickness in a PB bath make 40 &#181;m coronal sections.</li>
	<li>Wash extensively with 0.1 M PB. Follow the histological protocol for glial fibrillary acidic protein (GFAP) staining31.</li>
	<li>Mount sections on slides and cover them with a mounting medium containing 2-(4-amidinophenyl)-1H-indole-6-carboxamidine (DAPI) (see Table of Materials).</li>
</ol>
8. Confocal Microscopy
<ol>
	<li>Place the slides with the stained coronal sections under a 20x objective of a confocal microscope. Select the target region.</li>
	<li>Choose the optimal excitation and emission (exc/ems) filter sets for the dyes as follows: DAPI = 358/461 nm, DiI = 551/569 nm, and fluorescein (see Table of Materials) = 490/525 nm. 
	NOTE: Since staining varies per section, a proper range of minimal and maximal excitation and detection needs to be determined for each section, where the least dense and most dense regions both show emission.</li>
	<li>Choose the least dense region and set the laser intensity and detection to high values, and then verify at the densest regions whether these values cause oversaturation of detected emission. If so, lower the values and recheck them with the least dense region. Iterate these steps until arriving at the highest possible detection at low staining levels and proper, not oversaturated levels at highly stained areas. Repeat this process for all dyes.</li>
	<li>Use the tile scan function of the microscope with 512 x 512 pixels per tile to obtain a large overview of the probe insertion sites with an adequate resolution for post hoc processing.</li>
</ol>

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V. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Pharmacoresistant+epilepsy:+from+pathogenesis+to+current+and+emerging+therapies.">Pharmacoresistant epilepsy: from pathogenesis to current and emerging therapies.</a> Cleve Clinical Journal of Medicine. 77 (7), 457-467 (2010).</li></ol>

The authors have nothing to disclose.

By developing a new experimental protocol in an acute mouse model of epilepsy, SLEs could be successfully controlled with the help of a &#181;FIP implanted in the epileptic focus. Thanks to its capability to deliver GABA with temporal and spatial precision, 4AP-induced SLEs were controlled at the onset of the seizures. Treatment of epilepsy is theoretically possible if the control of the neural network discharges is achieved at the place of the seizure start. The presented protocol proved this possible if the localizatio...

Epilepsy is the fourth most common neurological disorder: about 1% of the population suffers from epilepsy, and about one-third of the affected have recurrent seizures. In most cases, seizures can be controlled with medication. However, drug treatment needs to be set for every patient individually, where proper dosing can take years to find1,2. Additionally, most of the medication has serious side effects that reduce the quality of life3,4,5,6,7. Finally, in 30% of the cases patients are resistant to medication, and in case of a constant single seizure generator locus, only resective neurosurgery can attenuate the occurrence of seizures8. Therefore, a major initiative in modern epilepsy research is to discover new strategies which can prevent recurrent seizures in patients at risk, while reducing the necessity of strong drug therapies and invasive resective surgeries.
Epileptic seizures occur when there is an imbalance within excitatory and inhibitory circuits either throughout the brain (generalized epilepsy) or in a localized part of the brain (focal epilepsy), such that neurons discharge in an abnormal fashion9,10,11. Antiepileptic drugs can act in two different ways in seizure prevention: either decreasing excitation or enhancing inhibition12. Specifically, they can either modify the electrical activity of neuronal cells by affecting ion channels in the cell membrane13 or act on chemical transmission between neurons by affecting the inhibitory neurotransmitter GABA or the excitatory glutamate in the synapses14,15. For some medications, the mode of action is unknown18. Also, drug treatments have a continuous effect on patients and cannot adapt to the prevalence dynamics of seizures. Ideally, drugs with specific mechanisms of action would act on the underlying epileptic processes. An optimal treatment would not touch the brain interictally but would act immediately when a seizure starts developing. In contrast to that, in all cases of epilepsy, medication now means a systematic treatment, affecting the whole brain and the whole body of the patient9.
Epileptic seizures can appear many years after the initial insult such as brain trauma. The period between the initial insult and the occurrence of the first spontaneous seizures is characterized by considerable molecular and cellular reorganizations, including neuronal death with the disappearance of neuronal network connections and axonal sprouting/neosynaptogenesis with the appearance of new connections19,20,21. Once seizures become recurrent, their frequency and severity tend to increase, involving more brain regions. It is important to distinguish the sites of seizure onset (epileptogenic regions) from propagation networks, as the rules of seizure genesis and propagation may differ. Research performed on human tissue and experimental models of epilepsy have provided important data regarding the reorganization of circuits and their ability to generate seizures20,21,22,23. However, it is difficult to determine if these reorganizations are adaptive responses or whether they are causally related to epileptogenesis or seizure genesis and propagation12.
Therefore, localizing the epileptic focus and applying antiepileptic drugs locally are one of the main challenges in contemporary epilepsy research. Several experiments using animal models of epilepsy and some clinical studies aimed to find the onset of the seizure events and define the underlying mechanisms in the brain24,25,26,27. To this end, we developed a new experimental protocol using the 4AP-induced epilepsy model28,29,30,31 in an acute mouse preparation, which allows the precise insertion of three devices into the given area of the hippocampus, where network activity in vivo is manipulated in a highly localized manner. Localized 4AP injection by a glass micropipette helps to induce epileptic SLEs in a localized spot in the hippocampus, while with the help of the novel polymer-based &#181;FIP probe the control of the seizure activity is achieved simultaneously by recording the neuronal electrical activity with the device&#8217;s recording sites. Hippocampal local field activity is also monitored with a multichannel silicon probe in a layer-specific manner in the cortex and in the hippocampus simultaneously.
The recently invented &#181;FIP probes work by using an applied electric field to push charged drugs stored in a microfluidic channel across an ion exchange membrane (IEM) and out to the surrounding tissue (Figure 1). The IEM selectively transports only one type of ion (cation or anion) and, thus, works to limit both passive diffusion in the &#8220;off&#8221; state and&#160; transport of oppositely charged species from the surrounding tissue into the device. The electric field is created on demand by applying a small voltage (&#60;1 V) between the source electrode which is internal to the microfluidic channel and a target electrode which is external to the device (in this case, the head screw on the animal model). The rate of drug delivery is proportional to the applied voltage and the measured current between the source and target electrodes. The precise tunability of drug delivery is one of the primary advantages of the &#181;FIP. Another critical advantage, compared to fluidic or pressure-based drug delivery systems, is that in the &#181;FIP there is only a negligible pressure increase at the drug delivery outlet as drugs are delivered across the IEM without their carrier solution.
There is a small amount of passive leaking of GABA when the &#181;FIP is &#8220;off&#8221;, but this was found not to effect SLEs. The &#181;FIP are custom-made following conventional microfabrication methods that we reported previously31.
Since one way of preventing recurrent seizures is the blockade of network discharges at the very beginning or even before the first seizure event, the presented method for delivering the inhibitory neurotransmitter GABA into the epileptic focus has great therapeutic potential for seizure control in patients with focal epilepsy. Since GABA is an endogenous substrate, it leaves intrinsic neuronal properties unchanged in physiological concentrations. The local application of low levels of GABA will only affect cells naturally responsive to inhibition, and will only cause similar effects to physiological inhibition, contrary to deep brain stimulation (DBS), which has unspecific actions by stimulating all cells of the neuronal network in its environment, causing a mixed response involving both excitation and inhibition. In conclusion, the proposed method provides a more specific approach to seizure control than DBS.

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr >
			<td >4AP</td>
			<td >Sigma</td>
			<td >275875</td>
			<td ></td>
		</tr>
		<tr >
			<td >Alexa Fluor 488</td>
			<td>Abcam</td>
			<td>ab15007</td>
			<td></td>
		</tr>
		<tr >
			<td >Amplifier</td>
			<td>Neuralynx, Montana, USA</td>
			<td >Digital Lynx 4SX</td>
			<td ></td>
		</tr>
		<tr >
			<td >Amplifier</td>
			<td >Ampliplex</td>
			<td >KJE-1001</td>
			<td ></td>
		</tr>
		<tr >
			<td >Atlas Stereotaxique&#160;</td>
			<td>Allen Atlas</td>
			<td>978-0470054086</td>
			<td></td>
		</tr>
		<tr >
			<td >Borosilica glass pipette</td>
			<td >Sutter</td>
			<td >BF120-69-15</td>
			<td ></td>
		</tr>
		<tr >
			<td >Brain Matrix</td>
			<td >WPI&#160;</td>
			<td >RBMA-200C</td>
			<td ></td>
		</tr>
		<tr >
			<td >Bone trimmer</td>
			<td >FST</td>
			<td >16109-14</td>
			<td ></td>
		</tr>
		<tr >
			<td >Confocal microscope</td>
			<td>Zeiss</td>
			<td>LSM 510</td>
			<td ></td>
		</tr>
		<tr >
			<td >Connector</td>
			<td>INSTECH</td>
			<td>SC20/15</td>
			<td ></td>
		</tr>
		<tr >
			<td >Coton tige</td>
			<td>Monoprix</td>
			<td>EMD 6107OD</td>
			<td></td>
		</tr>
		<tr >
			<td >Cover slip</td>
			<td >Menzel-Glass</td>
			<td >15747592</td>
			<td ></td>
		</tr>
		<tr >
			<td >DiI Stain&#160;</td>
			<td >Thermo Fisher</td>
			<td >D282</td>
			<td ></td>
		</tr>
		<tr >
			<td >DMSO</td>
			<td >Sigma</td>
			<td >11412-11</td>
			<td ></td>
		</tr>
		<tr >
			<td >Drill</td>
			<td>FOREDOM</td>
			<td>K1070</td>
			<td></td>
		</tr>
		<tr >
			<td >Forceps</td>
			<td >F.S.T.</td>
			<td >11412-11</td>
			<td ></td>
		</tr>
		<tr >
			<td >GABA</td>
			<td>Sigma</td>
			<td>A2129</td>
			<td></td>
		</tr>
		<tr >
			<td >GFAP Monoclonal Antibody</td>
			<td>Thermofisher</td>
			<td>53-9892-80</td>
			<td></td>
		</tr>
		<tr >
			<td >GOPS</td>
			<td>Sigma</td>
			<td>440167-100M</td>
			<td></td>
		</tr>
		<tr >
			<td >Hamilton seringe&#160;</td>
			<td>Hamilton&#160;</td>
			<td>80330</td>
			<td></td>
		</tr>
		<tr >
			<td >Headscrew</td>
			<td>Component Supply</td>
			<td>TX00-2FH</td>
			<td></td>
		</tr>
		<tr >
			<td >Heating pad&#160;</td>
			<td>Harvard apparatus</td>
			<td>341446</td>
			<td></td>
		</tr>
		<tr >
			<td >Injection Pump</td>
			<td>WPI&#160;</td>
			<td>UMP3-3</td>
			<td></td>
		</tr>
		<tr >
			<td >Keithley</td>
			<td >Tektoronix</td>
			<td >216A</td>
			<td ></td>
		</tr>
		<tr >
			<td >Ketamine</td>
			<td>Renaudin</td>
			<td>5787419</td>
			<td></td>
		</tr>
		<tr >
			<td >Magnetic holder</td>
			<td>Narishige</td>
			<td>GJ-1</td>
			<td></td>
		</tr>
		<tr >
			<td >Mice</td>
			<td>Charles River</td>
			<td>612</td>
			<td ></td>
		</tr>
		<tr >
			<td >Motoric manipulator</td>
			<td>Scientifica, UK</td>
			<td >IVM</td>
			<td ></td>
		</tr>
		<tr >
			<td >Na2HPO4</td>
			<td >Sigma</td>
			<td >255793</td>
			<td ></td>
		</tr>
		<tr >
			<td >NaH2PO4</td>
			<td >Sigma</td>
			<td >7558807</td>
			<td ></td>
		</tr>
		<tr >
			<td >NeuroTrace DiI&#160;</td>
			<td>Thermofisher</td>
			<td>N22880</td>
			<td></td>
		</tr>
		<tr >
			<td >Paper towel</td>
			<td>KIMBERLY CLARK</td>
			<td>7552000</td>
			<td></td>
		</tr>
		<tr >
			<td >PB</td>
			<td>Sigma</td>
			<td>P4417</td>
			<td></td>
		</tr>
		<tr >
			<td >PEDOT:PSS</td>
			<td>CLEVIOS</td>
			<td>81076212</td>
			<td></td>
		</tr>
		<tr >
			<td >PFA</td>
			<td>Acros Organic</td>
			<td>30525-89-4</td>
			<td></td>
		</tr>
		<tr >
			<td >Rectal temperature probe</td>
			<td>Harvard apparatus</td>
			<td>521591</td>
			<td></td>
		</tr>
		<tr >
			<td >Ropivacaine&#160;</td>
			<td>KABI</td>
			<td>1260216</td>
			<td></td>
		</tr>
		<tr >
			<td >Saline</td>
			<td>Sigma</td>
			<td>7982</td>
			<td></td>
		</tr>
		<tr >
			<td >Scalpel</td>
			<td>F.S.T</td>
			<td>AUST R195806</td>
			<td></td>
		</tr>
		<tr >
			<td >Seringue&#160;</td>
			<td>BD Medical</td>
			<td>324826</td>
			<td></td>
		</tr>
		<tr >
			<td >Serrefine clamp</td>
			<td>F.S.T</td>
			<td>18050-28</td>
			<td>4 is recommended</td>
		</tr>
		<tr >
			<td >Silicon probe</td>
			<td>NeuroNexus, Michigan, USA</td>
			<td>A2x16-10mm-50-500-177 or A1x16-5mm-150-703</td>
		</tr>
		<tr >
			<td >Stereotoxic frame</td>
			<td>Stoelting</td>
			<td>51733U</td>
			<td></td>
		</tr>
		<tr >
			<td >Superfrost Slide</td>
			<td >ThermoScientific</td>
			<td >J38000AMNZ</td>
			<td ></td>
		</tr>
		<tr >
			<td >Tubing</td>
			<td >INSTECH</td>
			<td >LS20</td>
			<td ></td>
		</tr>
		<tr >
			<td >Vaseline&#160;</td>
			<td>Laboratoire Gilbert</td>
			<td>3518646126611</td>
			<td></td>
		</tr>
		<tr >
			<td >Vectashield DAPI</td>
			<td>Vector Laboratories, California, USA</td>
			<td>H-1200-10</td>
			<td></td>
		</tr>
		<tr >
			<td >Vibratome, Leica VT1200S</td>
			<td>Leica Microsystems</td>
			<td>1491200S001</td>
			<td ></td>
		</tr>
		<tr >
			<td >Xylazine&#160;</td>
			<td>Bayer</td>
			<td>4007221032311</td>
			<td></td>
		</tr>
	</tbody>
</table>

electrophoretic delivery of &#x3b3;-aminobutyric acid (gaba) into epileptic focus prevents seizures in mice

Epilepsy is a group of neurological disorders which affects millions of people worldwide. Although treatment with medication is helpful in 70% of the cases, serious side effects affect the quality of life of patients. Moreover, a high percentage of epileptic patients are drug resistant; in their case, neurosurgery or neurostimulation are necessary. Therefore, the major goal of epilepsy research is to discover new therapies which are either capable of curing epilepsy without side effects or preventing recurrent seizures in drug-resistant patients. Neuroengineering provides new approaches by using novel strategies and technologies to find better solutions to cure epileptic patients at risk.
As a demonstration of a novel experimental protocol in an acute mouse model of epilepsy, a direct in situ electrophoretic drug delivery system is used. Namely, a neural probe incorporating a microfluidic ion pump (&#181;FIP) for on-demand drug delivery and simultaneous recording of local neural activity is implanted and demonstrated to be capable of controlling 4-aminopyridine-induced (4AP-induced) seizure-like event (SLE) activity. The &#947;-aminobutyric acid (GABA) concentration is kept in the physiological range by the precise control of GABA delivery to reach an antiepileptic effect in the seizure focus but not to cause overinhibition-induced rebound bursts. The method allows both the detection of pathological activity and intervention to stop seizures by delivering inhibitory neurotransmitters directly to the epileptic focus with precise spatiotemporal control.
As a result of the developments to the experimental method, SLEs can be induced in a highly localized manner that allows seizure control by the precisely tuned GABA delivery at the seizure onset.

Using the procedure presented here with a 4AP epilepsy model in anesthetized mice, control of epileptic seizures can be achieved in the epileptic focus. The precise localization of the implants (Figure 2) helped to record hippocampal local field potentials (LFPs, Figure 4), to induce small hippocampal seizures and to deliver GABA at the seizure onset. The localization of the implants was verified after each experiment by post hoc...

Watch this Scientific Journal Video about Electrophoretic Delivery of ÃŽÂ³-aminobutyric Acid GABA into Epileptic Focus Prevents Seizures in Mice at JoVE.com

Electrophoretic Delivery of &#947;-aminobutyric Acid GABA into Epileptic Focus Prevents Seizures in Mice

The challenge of epilepsy research is to develop novel treatments for patients where classical therapy is inadequate. Using a new protocol&#8212;with the help of an implantable drug delivery system&#8212;we are able to control seizures in anesthetized mice by the electrophoretic delivery of GABA into the epileptic focus.

Electrophoretic Delivery of &#x3B3;-aminobutyric Acid (GABA) into Epileptic Focus Prevents Seizures in Mice

Epilepsy is a group of neurological disorders which affects millions of people worldwide. Although treatment with medication is helpful in 70% of the ...

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8.9K Views.  Aix Marseille Université, Institut de Neurosciences des Systèmes (INS). The challenge of epilepsy research is to develop novel treatments for patients where classical therapy is inadequate. Using a new protocol—with the help of an implantable drug delivery system—we are able to control seizures in anesthetized mice by the electrophoretic delivery of GABA into the epileptic focus.

Video: Electrophoretic Delivery of γ-aminobutyric Acid GABA into Epileptic Focus Prevents Seizures in Mice

GABA-activated Single-channel and Tonic Currents in Rat Brain Slices

The GABAA channels are present in all neurons and are located both at synapses and outside of synapses where they generate phasic and tonic currents, respectively 4,5,6,7 The GABAA channel is a pentameric GABA-gated chloride channel. The channel subunits are grouped into 8 families (&alpha;1-6, &beta;1-3, &gamma;1-3, &delta;, &#949;, &theta;, &pi; and &rho;). Two alphas, two betas and one 3rd subunit form the functional channel 8. By combining studies of sub-type specific GABA-activated single-channel molecules with studies including all populations of GABAA channels in the neuron it becomes possible to understand the basic mechanism of neuronal inhibition and how it is modulated by pharmacological agents.
We use the patch-clamp technique 9,10 to study the functional properties of the GABAA channels in alive neurons in hippocampal brain slices and record the single-channel and whole-cell currents. We further examine how the channels are affected by different GABA concentrations, other drugs and intra and extracellular factors. For detailed theoretical and practical description of the patch-clamp method please see The Single-Channel Recordings edited by B Sakman and E Neher 10.

We use the patch-clamp technique to measure GABA-activated single-channel currents (GABAA channels, GABAA receptors) and the synaptic and tonic currents they generate in neurons. Activation of the channels decreases neuronal excitability in health and disease 1,2,3,4.

The GABAA channels are present in all neurons and are located both at synapses and outside of synapses where they generate phasic and tonic currents, ...

Efficient Gene Delivery into Multiple CNS Territories Using In Utero Electroporation

The ability to manipulate gene expression is the cornerstone of modern day experimental embryology, leading to the elucidation of multiple developmental pathways. Several powerful and well established transgenic technologies are available to manipulate gene expression levels in mouse, allowing for the generation of both loss- and gain-of-function models. However, the generation of mouse transgenics is both costly and time consuming. Alternative methods of gene manipulation have therefore been widely sought. In utero electroporation is a method of gene delivery into live mouse embryos1,2 that we have successfully adapted3,4. It is largely based on the success of in ovo electroporation technologies that are commonly used in chick5. Briefly, DNA is injected into the open ventricles of the developing brain and the application of an electrical current causes the formation of transient pores in cell membranes, allowing for the uptake of DNA into the cell. In our hands, embryos can be efficiently electroporated as early as embryonic day (E) 11.5, while the targeting of younger embryos would require an ultrasound-guided microinjection protocol, as previously described6. Conversely, E15.5 is the latest stage we can easily electroporate, due to the onset of parietal and frontal bone differentiation, which hampers microinjection into the brain. In contrast, the retina is accessible through the end of embryogenesis. Embryos can be collected at any time point throughout the embryonic or early postnatal period. Injection of a reporter construct facilitates the identification of transfected cells.
To date, in utero electroporation has been most widely used for the analysis of neocortical development1,2,3,4. More recent studies have targeted the embryonic retina7,8,9 and thalamus10,11,12. Here, we present a modified in utero electroporation protocol that can be easily adapted to target different domains of the embryonic CNS. We provide evidence that by using this technique, we can target the embryonic telencephalon, diencephalon and retina. Representative results are presented, first showing the use of this technique to introduce DNA expression constructs into the lateral ventricles, allowing us to monitor progenitor maturation, differentiation and migration in the embryonic telencephalon. We also show that this technique can be used to target DNA to the diencephalic territories surrounding the 3rd ventricle, allowing the migratory routes of differentiating neurons into diencephalic nuclei to be monitored. Finally, we show that the use of micromanipulators allows us to accurately introduce DNA constructs into small target areas, including the subretinal space, allowing us to analyse the effects of manipulating gene expression on retinal development.

Efficient Gene Delivery into Multiple CNS Territories Using In Utero Electroporation

In utero electroporation allows for rapid gene delivery in a spatially- and temporally-controlled manner in the developing central nervous system (CNS). Here we describe a highly adaptable in utero electroporation protocol that can be used to deliver expression constructs into multiple embryonic CNS domains, including the telencephalon, diencephalon and retina.

The ability to manipulate gene expression is the cornerstone of modern day experimental embryology, leading to the elucidation of multiple developmental ...

Extinction Training During the Reconsolidation Window Prevents Recovery of Fear

Fear is maladaptive when it persists long after circumstances have become safe. It is therefore crucial to develop an approach that persistently prevents the return of fear. Pavlovian fear-conditioning paradigms are commonly employed to create a controlled, novel fear association in the laboratory. After pairing an innocuous stimulus (conditioned stimulus, CS) with an aversive outcome (unconditioned stimulus, US) we can elicit a fear response (conditioned response, or CR) by presenting just the stimulus alone1,2 . Once fear is acquired, it can be diminished using extinction training, whereby the conditioned stimulus is repeatedly presented without the aversive outcome until fear is no longer expressed3. This inhibitory learning creates a new, safe representation for the CS, which competes for expression with the original fear memory4. Although extinction is effective at inhibiting fear, it is not permanent. Fear can spontaneously recover with the passage of time. Exposure to stress or returning to the context of initial learning can also cause fear to resurface3,4.
Our protocol addresses the transient nature of extinction by targeting the reconsolidation window to modify emotional memory in a more permanent manner. Ample evidence suggests that reactivating a consolidated memory returns it to a labile state, during which the memory is again susceptible to interference5-9. This window of opportunity appears to open shortly after reactivation and close approximately 6hrs later5,11,16, although this may vary depending on the strength and age of the memory15. By allowing new information to incorporate into the original memory trace, this memory may be updated as it reconsolidates10,11. Studies involving non-human animals have successfully blocked the expression of fear memory by introducing pharmacological manipulations within the reconsolidation window, however, most agents used are either toxic to humans or show equivocal effects when used in human studies12-14. Our protocol addresses these challenges by offering an effective, yet non-invasive, behavioral manipulation that is safe for humans.
By prompting fear memory retrieval prior to extinction, we essentially trigger the reconsolidation process, allowing new safety information (i.e., extinction) to be incorporated while the fear memory is still susceptible to interference. A recent study employing this behavioral manipulation in rats has successfully blocked fear memory using these temporal parameters11. Additional studies in humans have demonstrated that introducing new information after the retrieval of previously consolidated motor16, episodic17, or declarative18 memories leads to interference with the original memory trace14. We outline below a novel protocol used to block fear recovery in humans.

Conditioned fear can be diminished through an inhibitory process called extinction, but can resurface under conditions such as the passage of time or exposure to stress. Our protocol presents a novel way of preventing fear recovery by introducing extinction during the reconsolidation window (the re-storage phase of a reactivated memory).

Fear is maladaptive when it persists long after circumstances have become safe. It is therefore crucial to develop an approach that persistently prevents ...

Direct Intraventricular Delivery of Drugs to the Rodent Central Nervous System

Due to an inability to cross the blood brain barrier, certain drugs need to be directly delivered into the central nervous system (CNS). Our lab focuses specifically on antisense oligonucleotides (ASOs), though the techniques shown in the video here can also be used to deliver a plethora of other drugs to the CNS. Antisense oligonucleotides (ASOs) have the capability to knockdown sequence-specific targets 1 as well as shift isoform ratios of specific genes 2. To achieve widespread gene knockdown or splicing in the CNS of mice, the ASOs can be delivered into the brain using two separate routes of administration, both of which we demonstrate in the video.
The first uses Alzet osmotic pumps, connected to a catheter that is surgically implanted into the lateral ventricle. This allows the ASOs to be continuously infused into the CNS for a designated period of time. The second involves a single bolus injection of a high concentration of ASO into the right lateral ventricle. Both methods use the mouse cerebral ventricular system to deliver the ASO to the entire brain and spinal cord, though depending on the needs of the study, one method may be preferred over the other.

We describe a method to target drugs to the central nervous system by either implanting a catheter or performing a bolus injection into the right lateral ventricle in mice. We focus specifically on the delivery of antisense oligonucleotides. This technique is readily adaptable to other drugs and to rats.

Due to an inability to cross the blood brain barrier, certain drugs need to be directly delivered into the central nervous system (CNS). Our lab focuses ...

Fast Micro-iontophoresis of Glutamate and GABA: A Useful Tool to Investigate Synaptic Integration

One of the fundamental interests in neuroscience is to understand the integration of excitatory and inhibitory inputs along the very complex structure of the dendritic tree, which eventually leads to neuronal output of action potentials at the axon. The influence of diverse spatial and temporal parameters of specific synaptic input on neuronal output is currently under investigation, e.g. the distance-dependent attenuation of dendritic inputs, the location-dependent interaction of spatially segregated inputs, the influence of GABAergig inhibition on excitatory integration, linear and non-linear integration modes, and many more.
With fast micro-iontophoresis of glutamate and GABA it is possible to precisely investigate the spatial and temporal integration of glutamatergic excitation and GABAergic inhibition. Critical technical requirements are either a triggered fluorescent lamp, light-emitting diode (LED), or a two-photon scanning microscope to visualize dendritic branches without introducing significant photo-damage of the tissue. Furthermore, it is very important to have a micro-iontophoresis amplifier that allows for fast capacitance compensation of high resistance pipettes. Another crucial point is that no transmitter is involuntarily released by the pipette during the experiment.
Once established, this technique will give reliable and reproducible signals with a high neurotransmitter and location specificity. Compared to glutamate and GABA uncaging, fast iontophoresis allows using both transmitters at the same time but at very distant locations without limitation to the field of view. There are also advantages compared to focal electrical stimulation of axons: with micro-iontophoresis the location of the input site is definitely known and it is sure that only the neurotransmitter of interest is released. However it has to be considered that with micro-iontophoresis only the postsynapse is activated and presynaptic aspects of neurotransmitter release are not resolved. In this article we demonstrate how to set up micro-iontophoresis in brain slice experiments.

In this article we introduce fast micro-iontophoresis of neurotransmitters as a technique to investigate integration of postsynaptic signals with high spatial and temporal precision.

One of the fundamental interests in neuroscience is to understand the integration of excitatory and inhibitory inputs along the very complex structure of ...

Lineage-reprogramming of Pericyte-derived Cells of the Adult Human Brain into Induced Neurons

Direct lineage-reprogramming of non-neuronal cells into induced neurons (iNs) may provide insights into the molecular mechanisms underlying neurogenesis and enable new strategies for in vitro modeling or repairing the diseased brain. Identifying brain-resident non-neuronal cell types amenable to direct conversion into iNs might allow for launching such an approach in situ, i.e. within the damaged brain tissue. Here we describe a protocol developed in the attempt of identifying cells derived from the adult human brain that fulfill this premise. This protocol involves: (1) the culturing of human cells from the cerebral cortex obtained from adult human brain biopsies; (2) the in vitro expansion (approximately requiring 2-4 weeks) and characterization of the culture by immunocytochemistry and flow cytometry; (3) the enrichment by fluorescence-activated cell sorting (FACS) using anti-PDGF receptor-&#946; and anti-CD146 antibodies; (4) the retrovirus-mediated transduction with the neurogenic transcription factors sox2 and ascl1; (5) and finally the characterization of the resultant pericyte-derived induced neurons (PdiNs) by immunocytochemistry (14 days to 8 weeks following retroviral transduction). At this stage, iNs can be probed for their electrical properties by patch-clamp recording. This protocol provides a highly reproducible procedure for the in vitro lineage conversion of brain-resident pericytes into functional human iNs.

Targeting brain-resident cells for direct lineage-reprogramming offers new perspectives for brain repair. Here we describe a protocol of how to prepare cultures enriched for brain-resident pericytes from the adult human cerebral cortex and convert these into induced neurons by retrovirus-mediated expression of the transcription factors Sox2 and Ascl1.

Direct lineage-reprogramming of non-neuronal cells into induced neurons (iNs) may provide insights into the molecular mechanisms underlying neurogenesis ...

Subpial Adeno-associated Virus 9 (AAV9) Vector Delivery in Adult Mice

The successful development of a subpial adeno-associated virus 9 (AAV9) vector delivery technique in adult rats and pigs has been reported on previously. Using subpially-placed polyethylene catheters (PE-10 or PE-5) for AAV9 delivery, potent transgene expression through the spinal parenchyma (white and gray matter) in subpially-injected spinal segments has been demonstrated. Because of the wide range of transgenic mouse models of neurodegenerative diseases, there is a strong desire for the development of a potent central nervous system (CNS)-targeted vector delivery technique in adult mice. Accordingly, the present study describes the development of a spinal subpial vector delivery device and technique to permit safe and effective spinal AAV9 delivery in adult C57BL/6J mice. In spinally immobilized and anesthetized mice, the pia mater (cervical 1 and lumbar 1-2 spinal segmental level) was incised with a sharp 34 G needle using an XYZ manipulator. A second XYZ manipulator was then used to advance a blunt 36G needle into the lumbar and/or cervical subpial space. The AAV9 vector (3-5 &#181;L; 1.2 x 1013 genome copies (gc)) encoding green fluorescent protein (GFP) was then injected subpially. After injections, neurological function (motor and sensory) was assessed periodically, and animals were perfusion-fixed 14 days after AAV9 delivery with 4% paraformaldehyde. Analysis of horizontal or transverse spinal cord sections showed transgene expression throughout the entire spinal cord, in both gray and white matter. In addition, intense retrogradely-mediated GFP expression was seen in the descending motor axons and neurons in the motor cortex, nucleus ruber, and formatio reticularis. No neurological dysfunction was noted in any animals. These data show that the subpial vector delivery technique can successfully be used in adult mice, without causing procedure-related spinal cord injury, and is associated with highly potent transgene expression throughout the spinal neuraxis.

Subpial Adeno-associated Virus 9 AAV9 Vector Delivery in Adult Mice

The goal of the present study was to develop and validate the potency and safety of spinal adeno-associated virus 9 (AAV9)-mediated gene delivery by using a novel subpial gene delivery technique in adult mice.

The successful development of a subpial adeno-associated virus 9 (AAV9) vector delivery technique in adult rats and pigs has been reported on previously. ...

Whole-cell Currents Induced by Puff Application of GABA in Brain Slices

Pharmacological administration is commonly used when conducting whole-cell patch-clamp recording in brain slices. One of the best methods of drug application during electrophysiological recording is the puff technique, which can be used to study the effect of pharmacological reagents on neuronal activities in brain slices. The greatest advantage of puff application is that the drug concentration around the recording site increases rapidly, thus preventing desensitization of membrane receptors. Successful use of puff application involves careful attention to the following elements: the concentration of the drug, the parameters of the puff micropipette, the distance between the tip of a puff micropipette and the neuron recorded, and the duration and pressure driving the puff (pounds per square inch, psi). This article describes a step-by-step procedure for recording whole-cell currents induced by puffing gamma-aminobutyric acid (GABA) onto a neuron of a prefrontal cortical slice. Notably, the same procedure can be applied with minor modifications to other brain areas such as the hippocampus and the striatum, and to different preparations, such as cell cultures.

We describe the puff technique, by which pharmacological reagents can be administered during whole-cell patch-clamp recording, and highlight various aspects of the features that are crucial for its success.

Pharmacological administration is commonly used when conducting whole-cell patch-clamp recording in brain slices. One of the best methods of drug ...

Delivery of Antibodies into the Murine Brain via Convection-enhanced Delivery

Convection-enhanced delivery (CED) is a neurosurgical technique enabling effective perfusion of large brain volumes using a catheter system. Such an approach provides a safe delivery method by-passing the blood brain barrier (BBB), thus allowing treatment with therapeutics with poor BBB-permeability or those for which systemic exposure is not desired, e.g., due to toxicity. CED requires optimization of the catheter design, injection protocol, and properties of the infusate. With this protocol we describe how to perform CED of a solution containing up to 20 &#181;g of an antibody into the caudate putamen of mice. It describes preparation of step catheters, testing them in vitro and performing the CED in mice using a ramping injection program. The protocol can be readily adjusted for other infusion volumes and can be used for injecting various tracers or pharmacologically active or inactive substances, including chemotherapeutics, cytokines, viral particles, and liposomes.

Convection-enhanced delivery (CED) is a method enabling effective delivery of therapeutics into the brain by direct perfusion of large tissue volumes. The procedure requires the use of catheters and an optimized injection procedure. This protocol describes a methodology for CED of an antibody into a mouse brain.

Convection-enhanced delivery (CED) is a neurosurgical technique enabling effective perfusion of large brain volumes using a catheter system. Such an ...

Delivery of Antibodies into the Brain Using Focused Scanning Ultrasound

Only a small fraction of therapeutic antibodies targeting brain diseases are taken up by the brain. Focused ultrasound offers a possibility to increase uptake of antibodies and engagement through transient opening of the blood-brain barrier (BBB). In our laboratory, we are developing therapeutic approaches for neurodegenerative diseases in which an antibody in various formats is delivered across the BBB using microbubbles, concomitant with focused ultrasound application through the skull targeting multiple spots, an approach we refer to as scanning ultrasound (SUS). The mechanical effects of microbubbles and ultrasound on blood vessels increases paracellular transport across the BBB by transiently separating tight junctions and enhances vesicle- mediated transcytosis, allowing antibodies and therapeutic agents to effectively cross. Moreover, ultrasound also facilitates the uptake of antibodies from the interstitial brain into brain cells such as neurons where the antibody distributes throughout the cell body and even into neuritic processes. In our studies, fluorescently labeled antibodies are prepared, mixed with in-house prepared lipid-based microbubbles and injected into mice immediately before SUS is applied to the brain. The increased antibody concentration in the brain is then quantified. To account for alterations in normal brain homeostasis, microglial phagocytosis can be used as a cellular marker. The generated data suggest that ultrasound delivery of antibodies is an attractive approach to treat neurodegenerative diseases.

Presented here is a protocol to transiently open the blood-brain barrier (BBB) either focally or throughout a mouse brain to deliver fluorescently-labeled antibodies and activate microglia. Also presented is a method to detect the delivery of antibodies and microglia activation by histology.

Only a small fraction of therapeutic antibodies targeting brain diseases are taken up by the brain. Focused ultrasound offers a possibility to increase ...