Supported by the National Natural Science Foundation of China Grants 31522029, 31770929 and 31371149 (to Haitao Wu), Program 973 (2014CB542203) from the State Key Development Program for Basic Research of China (to Haitao Wu), and Grant Z161100000216154 from the Beijing Municipal Science and Technology Commission (to Haitao Wu). The authors thank all the members of the Haitao Wu laboratory for their encouragement and discussions. The authors are extremely grateful to Zhenwei Liu for his help with debugging the apparatus.

Animal experimental procedures followed the institutional guidelines of the Beijing Institute of Basic Medical Sciences (Beijing, China) and the Chinese governmental regulations for the Care and Use of Laboratory Animals. The mice (adult male, 26 ~ 30 g) were housed and kept at a constant temperature of 23 &#176;C, with water and food ad libitum, under a 12-h light/12-h dark cycle (lights on at 7:00 a.m.). All experimental procedures were performed during the light cycle.
1. Surgical Preparation
NOTE: A custom electrode was homemade using the method modified from Halpern&#39;s report23.
<ol>
	<li>Use 2 copper micro-wires (with an outer diameter of 200 &#181;m) as the parallel bipolar electrode. Wrap the electrodes around each other to form cylindrical electrodes for the stimulation. 
	NOTE: Here, the electrode length within either the PP or DG was varied according to the anatomical depth of the nucleus.</li>
	<li>Sterilize the electrodes using an Ethylene oxide disinfection cabinet. Handle and store the homemade 2-channel micro-wire electrodes without any contamination. The electrodes should be strictly sterilized before any surgical implantation.</li>
	<li>Autoclave the surgical instruments.</li>
	<li>Before the surgery, inject the mouse with pentobarbital (20 - 50 mg/kg) intraperitoneally, to achieve a surgical plane of anesthesia.</li>
	<li>Inject the mouse with a sterile penicillin G procaine suspension (75,000 U, I.M.) and an analgesic agent (carprofen, 0.05 - 1&#160;mg/kg, SC) to decrease the risk of infections and pain, respectively24.</li>
	<li>After the animal anesthesia, apply an eye lubricant to both eyes to prevent eye dryness. Ensure the induction of the sedation and the depth of the anesthesia by the animal&#39;s reflexes in response to toe pinching and proceed with the operation only when no response occurs. 
	NOTE: Vet ointment is another option to prevent eye dryness.</li>
	<li>Shave most of the fur on the top of the mouse head with electrical hair clippers or a razor blade, sterilize the shaved area with three alternating scrubs of 70% alcohol and betadine solution and proceed with the operation after the betadine has dried on the skin.</li>
</ol>
2. High-Frequency Stimulation Surgery25,26
NOTE: In this step, an electrode is unilaterally implanted into the dorsal DG or medial PP area, a part of the hippocampus with critical roles in episodic learning and memory.
<ol>
	<li>Place the anesthetized mouse onto the stereotactic frame. To keep the mouse&#39;s head immobile, align the ear bars properly and install them quickly. Gently tighten the ear bars so that they support the centered head and secure the mouse slightly.</li>
	<li>Use a thermostatic pad to maintain the animal&#39;s body temperature at 37 &#177; 0.5 &#176;C.</li>
	<li>Use forceps to grasp the skin anterior to the mouse ears and cut it with sterilized scissors to remove about 1-cm2 area of skin on the top of the mouse&#39;s skull. 
	NOTE: A small amount of bleeding could be observed along the edges of the cut skin.</li>
	<li>Remove the muscle and other attachment overlying the skull with a cotton swab to expose the skull surface. Use a cotton swab and dissecting chisel to clean all periosteum, tendons and connective tissue from the skull&#8217;s surface completely to identify the sagittal and lambda sutures lines.</li>
	<li>Identify the bregma and lambda point. The intersection of the coronal suture and the sagittal suture on the superior middle portion of the calvaria is where the bregma is located. The intersection of the lambdoid suture by the sagittal suture is where the lambda is visible. Use the bregma as the original point to set the other nucleus position.</li>
	<li>After the identification of the bregma and lambda position on the skull, adjust these two points in the horizontal level. Make sure that the height of these two points is almost the same in the dorsal-ventral direction (any errors should be &#60; 0.05 mm). Also, make sure that the skull is level in the medial-lateral direction in the same way26.</li>
	<li>For a unilateral surgery, mount a homemade 2-channel copper micro-wire electrode into the rotating electrode holder on the stereotactic surgical frame.</li>
	<li>Place the electrode vertically above the bregma point at first, and set it as original site. Use the digital displayed stereotactic surgical frame to indicate the anterior-posterior (AP), medial-lateral (ML), and dorso-ventral (DV) positions. 
	NOTE: To prevent the electrode from getting bent, do not touch the electrode tip onto the skull during the operation.</li>
	<li>Move the electrode gently to the correct position (Figure 1A, AP-2.1/ML &#177; 1.0 for DG and AP-3.8/ML &#177; 3.0 for PP, respectively)27 above the skull without any touch. When the desired site was identified for electrode insertion, use the dorsoventral stereotactic adjustments to higher the electrodes holder, and mark the position with a black marker. 
	NOTE: The exact location in reference to the stereotaxic coordinates may vary by mouse strain, weight, and sex. Adult mice of the same sex should be used to minimize any variation in the location. If possible, pre-operational anatomical scans should be used to identify the location on an individual subject basis27.</li>
	<li>Use a hand-held micromanipulator-assisted drill (burr size = 0.5 mm) to make burr holes at the marked position precisely. Sterilize the tip of the burr with 70% ethanol before drilling. Move up the mini drill bit through the skull along the Z-axis 0.8 - 1.0 mm without changing its X-Y position. Use a sterile cotton to stop any bleeding during the operation. 
	NOTE: To avoid over-drilling and excessive heat generation during the drilling, ensure the drilling speed is 15,000 - 18,000 revolutions per minute (RPM), and frequently withdraw the drill bit from burr hole every 8 - 10 s.</li>
	<li>Keep drilling until the dura is exposed. Use a bent needle to remove any bone scraps that are generated during the drilling and make a pinhole on the dura without damaging the underlying soft brain tissue. 
	NOTE: To avoid damaging the brain tissue, the bent blunt needle can also be replaced with a fine blunt forceps.</li>
	<li>To confirm that the burr holes have been made properly at the desired position, reduce the electrode height by an adjustment of the stereotactic surgery frame and ensure that the electrode can be inserted smoothly through the burr hole without touching any obstacles. If so, slowly insert the electrodes to a depth of 1.8 mm for the DG (or 3.0 mm for the PP) from the surface of the skull located at the medial PP to the DG (PP-DG). 
	NOTE: Wipe off any cerebrospinal fluid or blood outflows from the burr holes with sterile cotton during the process of the electrode implantation.</li>
	<li>After inserting the micro-wire electrode into the skull hole, hold it in place with adequate dental cement using a small spatula. As the cement thickens, mold it around the inserted electrode and screws to form a nice smooth cap. Debride and disinfect the wound edges.</li>
	<li>Disengage the electrode from the electrode holder. Remove the mouse from the stereotactic frame and return it to a warm cage. 
	NOTE: After surgery, wait with returning the animals to their cages until they have fully recovered from the anesthesia with sufficient consciousness.</li>
	<li>Let the mouse move freely and allow it to recover for 2 days in the cage. Then, connect the electrode implanted in the mouse brain to a programmable stimulator via leads. Set up the software interface as presented in Figure 2. Set the HFS parameters as 6 series of 6 trains of 6 pulses at 400 Hz (Figure 1B, 100 ms between the trains, 20 s between the series)28. Set the stimulation intensity to 200 &#956;A.</li>
	<li>Deliver an HFS for the desired period of time as set up. Deliver the stimulation 2x a day at 8:00 a.m. and at 8:00 p.m.</li>
	<li>For the control group, implant electrodes in the mice and do not perform a stimulation.</li>
	<li>For the c-fos staining, stimulate the experimental mice acutely (2x per day). For the BrdU labeling, divide the experimental mice into an acute stimulation group (applied 2x per day) and a chronic stimulation group (applied 10x in 5 days, 2x per day). During the course of the stimulation, make sure the mouse is awake. Remember to ensure the leads are not twisted. 
	NOTE: At the last day of the procedure, the mice in the acute stimulation group were stimulated simultaneously.</li>
	<li>When the stimulation is done, carefully disconnect the electrode pin from the electrode holder and the connector of the recording system.</li>
</ol>
3. Immunofluorescence Staining and Bromodeoxyuridine Labeling29
<ol>
	<li>For the c-fos and Notch1 staining, about 3 h after the last HF-DBS stimulation, anesthetize the mice by injecting pentobarbital (20 - 50 mg/kg, I.P.) to achieve a surgical plane of anesthesia.</li>
	<li>For the BrdU labeling, about 12 h after the last HF-DBS stimulation, give 6 injections of BrdU to the control and the stimulated mice at 2-h intervals (50 mg/kg in 0.9% saline, I.P.). 36 h after the last injection, anesthetize the mice by injecting pentobarbital (20 - 50 mg/kg, I.P.) to achieve a surgical plane.</li>
	<li>Make incisions using a scalpel through the rib cage to expose the heart and then pass a 22-gauge blunt perfusion needle to the left ventricle. Make a small incision in the right atrium. Transcardially perfuse it with 50 mL of cold phosphate-buffered saline (PBS) followed by 50 mL of cold 4% paraformaldehyde (PFA) in 1x PBS (pH 7.4). 
	NOTE: Switch the PBS to the PFA when the mouse blood in vivo is replaced and the fluid runs clear. A good perfusion can be judged by the clearing of the liver. The perfusion should be stopped once fixation tremors have been observed30.</li>
	<li>Decapitate the mouse with ophthalmic scissors, make an incision on the skin and expose the skull. Using forceps, chip off the skull slightly and carefully dissect the whole mouse brain. Post-&#64257;x the brain in 4% paraformaldehyde for an additional 2 days and transfer it to a 30% sucrose solution at 4 &#176;C overnight.</li>
	<li>Using a cryostat, cut the brains into 20-&#181;m coronal sections. Transfer and mount the sections to the glass slide with a brush and store them at -20 &#176;C. 
	NOTE: Animals with electrode misplacements or excessive mechanical damage will be excluded from the subsequent analysis.</li>
	<li>Transfer the slides into normal PBS (0.1 M, pH 7.4) containing no fixative. Wash the slices with PBS for 3 - 4x for 5 min each to remove any excess fixative.For the BrdU-labeled group, incubate the sections in 2-N HCl for 30 min at 37 &#176;C, and rinse them in a 0.1-M borate buffer (pH 8.4; 10 min). Then wash the slices 2x for 5 min in PBS. 
	NOTE: Immunostaining is then performed.</li>
	<li>Treat slices by incubation in blocking solution (1 % normal goat serum, 3 % bovine serum albumin &#60;BSA&#62; , 0.5 % Triton X-100 and 0.02 % sodium azide in 1 x PBS) for 1 h at room temperature (RT).</li>
	<li>Incubate the slices with anti-c-fos polyclonal antibody (1:500; rabbit), anti-Notch1 antibody (1:50; goat), or anti-BrdU antibody (1:800; rat) in the blocking solution at 4 &#176;C overnight.</li>
	<li>Wash the slices 3x in PBS, then incubate them with Alexa Fluor 568-conjugated anti-rabbit (1:500), Alexa Fluor 488-conjugated anti-goat (1:1,000), or Alexa Fluor 488-conjugated anti-rat (1:500) secondary antibody for 1 h at RT.</li>
	<li>Wash the samples 3x in PBS (10 min each time). Cover the immuno-labeled brain sections using an anti-fade reagent mounting medium containing DAPI. Let the slices air-dry and store them protected from light at 4 &#176;C.</li>
	<li>Acquire images of the hippocampal areas from the stimulated and unstimulated sides of the brain with a high-resolution multi-channel (sequential) scanning confocal microscope. Set the imaging parameters consistently between the control and experimental groups.</li>
	<li>Perform a mosaic imaging using a high-magnification objective (20X air objective, NA 0.45) to acquire continuous 3D (XYZ) images of adjacent fields of view using the motorized stage and utilize appropriate software to stitch them together to make large composite images. 
	NOTE: Tiling functions include true stitching and smoothing options for improved seamless images. Composite images are quickly and easily prepared using the stitching function, to form an image over a wide area.</li>
	<li>For c-fos and Notch1 staining, perform a positive nuclei quantitative analysis of the immunofluorescent density21 of the stitched images. Randomly select 3 areas in the rostral, dorsal, and ventral hippocampal DG subregions and measure the immunofluorescent density in a double-blinded manner using Image J software (http://rsb.info.nih.gov/ij/)31.</li>
	<li>For a BrdU labeling, analyze the slices around the drilling point through the dorsal hippocampus (-1.7 mm to -2.7 mm relative to the bregma) in a double-blinded manner.</li>
	<li>Count only BrdU-positive neurons in the DG. Include the cells lying within 2-cell diameters of the granule cell in the count. 
	NOTE: Here, counting was performed using a 40X air objective. Randomly, 3 - 5 sections per animal were counted, and the counts were averaged and expressed as means per DG19.</li>
</ol>

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(94), e52220 (2014).</li><li>Pizzolato, G., Mandat, T. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Deep+brain+stimulation+for+movement+disorders.">Deep brain stimulation for movement disorders.</a> Frontiers in Integrative Neuroscience. 6, 2 (2012).</li><li>Tierney, T. S., Sankar, T., Lozano, A. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Some+recent+trends+and+further+promising+directions+in+functional+neurosurgery.">Some recent trends and further promising directions in functional neurosurgery.</a> Acta Neurochirurgica Supplement. 117 (117), 87-92 (2013).</li><li>Laxton, A. W., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+phase+I+trial+of+deep+brain+stimulation+of+memory+circuits+in+Alzheimer's+disease.">A phase I trial of deep brain stimulation of memory circuits in Alzheimer's disease.</a> Annals of Neurology. 68 (4), 521-534 (2010).</li><li>Min, H. K., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Deep+brain+stimulation+induces+BOLD+activation+in+motor+and+non-motor+networks:+an+fMRI+comparison+study+of+STN+and+EN/GPi+DBS+in+large+animals.">Deep brain stimulation induces BOLD activation in motor and non-motor networks: an fMRI comparison study of STN and EN/GPi DBS in large animals.</a> NeuroImage. 63 (3), 1408-1420 (2012).</li><li>Kukurba, K. R., Montgomery, S. B. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=.">.</a> RNA sequencing and analysis. 2015 (11), 951-969 (2015).</li><li>Kawashima, T., Okuno, H., Bito, H. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+new+era+for+functional+labeling+of+neurons:+activity-dependent+promoters+have+come+of+age.">A new era for functional labeling of neurons: activity-dependent promoters have come of age.</a> Frontiers in Neural Circuits. 8, 37 (2014).</li><li>Liu, J., Solway, K., Messing, R. O., Sharp, F. R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Increased+neurogenesis+in+the+dentate+gyrus+after+transient+global+ischemia+in+gerbils.">Increased neurogenesis in the dentate gyrus after transient global ischemia in gerbils.</a> Journal of Neuroscience. 18 (19), 7768-7778 (1998).</li><li>Kuhn, H. G., Dickinson-Anson, H., Gage, F. H. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Neurogenesis+in+the+dentate+gyrus+of+the+adult+rat:+age-related+decrease+of+neuronal+progenitor+proliferation.">Neurogenesis in the dentate gyrus of the adult rat: age-related decrease of neuronal progenitor proliferation.</a> Journal of Neuroscience. 16 (6), 2027-2033 (1996).</li></ol>

The authors have nothing to declare.

The HF-DBS technique has been widely used as a powerful tool for the treatment of many neurological disorders since the 1990s. So far, the landmark work of HF-DBS is for the treatment of Parkinson&#39;s disease and essential tremor, which has attracted much attention and interest both in the clinic and scientific community. There are various types of ongoing HF-DBS studies by many groups for HF-DBS&#39;s therapeutic application in certain neurological and psychiatric disorders32,<sup cl...

HF-DBS is a neurosurgical technology for electrical stimulation in the brain, which has been developed since the 1870s1. In the late 1980s, HFS was first used as a potential therapeutic intervention for Parkinson&#39;s disease and other movement disorders2. In the past few decades, HF-DBS has been more and more widely used in the treatment of brain disorders which are currently untreatable by a traditional therapeutic strategy. Particularly, due to the accuracy improvement of the HFS electrode, the highly effective outcomes, and minimal side effects, the number of brain disorders treated by HF-DBS has significantly increased over the past decades3,4,5. For example, HF-DBS has been approved by the US Food and Drug Administration (FDA) for the treatment of Parkinson&#39;s disease (PD), Alzheimer&#39;s type dementia, essential tremor, and other types of movement disorders2,6,7. In PD patients, the dopaminergic medication is reduced up to 50% during HF-DBS8. In addition to the successful treatment of movement disorders, HF-DBS has also demonstrated its powerful effects in the treatment of psychiatric diseases in the clinic, and for cognitive augmentation as well2,9,10,11. It should be noted that the research of HFS for the treatment of other psychiatric disorders are in various stages, offering much promise to patients12.
Although many studies have demonstrated that a focal HFS has both local and remote effects throughout the brain13, the neurological and molecular mechanisms of the effects remain elusive2,14. In the clinic, therapeutic HF-DBS is usually applied in a long-term manner for the treatment of Parkinson&#39;s disease and chronic pain, etc. Many opinions are raised to explain the improvement generated by an HF-DBS treatment, among which one possibility that the HFS current modulates the neuronal network activity, probably by a repetitive depolarization of the axons in the vicinity of the implanted HFS electrode. Or, HF-DBS may change the discharge rate of the output neurons and the projected targets. Also, HF-DBS may lead to long-term synaptic changes, including long-term potentiation (LTP) and long-term depression (LTD), which may contribute to a symptomatic improvement. So far, it is still unclear whether HFS in&#64258;uences the key molecular events that regulate cellular processes such as adult neurogenesis in vivo. Several lines of studies have demonstrated that HFS in rodents could mimic similar neural responses of clinically applied DBS15,16. To understand the underlying cellular mechanisms of HF-DBS, in this study, we first set up an in vivo HFS methodology in mice in an acute (one day) or chronic (five days) manner. Secondly, we set up an activation analysis methodology to determine the alteration of the neuronal activity and neurogenesis after an HF-DBS delivery.
Given that the neuronal production from neural stem cells is abundant during the embryonic development but continues throughout adult life, the hippocampal subgranular zone is one of the major areas where the neurogenesis occurs. The process of neurogenesis is influenced by many physiological and pathological factors. In certain epileptic cases, the hippocampal neurogenesis is dramatically decreased17,18. In addition, a single electroconvulsive therapy could significantly increase the neuronal production in the dentate gyrus19. These observations suggest that the electrophysiological activity plays a critical role in the regulation of adult neurogenesis and synaptic plasticity in hippocampal neurons. Therefore, to further demonstrate the effects of HF-DBS on neuronal activity and neurogenesis, we first carry out an immunostaining assay of the immediate early gene (IEG) c-fos which is a well-known marker of short-term neuronal activity resulting from experience20. Notch1 signaling is also detected to monitor the signaling activation after the HFS delivery21,22. Moreover, we also detect the neuronal production by a BrdU labeling analysis after the HF-DBS induction in various manners, though BrdU staining can also be a marker for gliogenesis.
In the present study, two HFS methodologies are adapted to target the activation of the hippocampal DG directly and indirectly. The electrode is implanted into the DG directly or implanted into the medial perforant path (PP) which sends projections to activate the DG neurons. For the HF-DBS induction, a programmable stimulator is presented for a continuous stimulation via the fixed electrode onto the mouse head. To determine the effects of HFS on neuronal activation and neurogenesis, we detect the expression of c-fos and Notch1 by immunofluorescent staining and the number of BrdU-incorporated positive neurons in the hippocampal DG region, respectively, after the HFS treatment. Particularly, the effects of the HF-DBS on the neurogenesis in the DG are compared between an acute and a chronic stimulation manner, or between a direct and an indirect stimulation manner, respectively.

<table><tbody><tr><td>Brain stereotaxic instrument</td><td>Stoelting</td><td>51730D</td><td>Stereotactic intracranial implantation for mouse</td></tr><tr><td>Stimulator</td><td>A-M systems</td><td>Model 3800</td><td>MultiStim 8-Channel programmable stimulator</td></tr><tr><td>Dental driller</td><td>Saeshin Precision Co., Ltd</td><td>STRONG 90</td><td>For drilling and crainiotomy&amp;nbsp;</td></tr><tr><td>Burr</td><td>Meisinger</td><td>HM1 005#</td><td>For drilling and crainiotomy&amp;nbsp;</td></tr><tr><td>Digidata 1550 Digitizer</td><td>Molecular Devices</td><td>AXON 1550</td><td>High-resolution data acquisition</td></tr><tr><td>Cryotome</td><td>Thermo Fisher Scientific</td><td>Thermo Cryotome FSE</td><td>Cutting frozen sections of specimens</td></tr><tr><td>Confocal microscope</td><td>Olympus</td><td>FV-1200</td><td>Japan, with 20x Objective (NA 0.45)</td></tr><tr><td>Mouse surgery tools</td><td>F.S.T.</td><td>14084-08,11254-20,16109-14</td><td>Scissors, forceps, bone cutter, holders etc.</td></tr><tr><td>Pentobarbital sodium</td><td>R&amp;amp;D systems</td><td>4579</td><td>20-50mg/kg for i.p. injection</td></tr><tr><td>Penicillin G&amp;nbsp;</td><td>Sigma-Aldrich</td><td>P3032</td><td>75,000 U for i.m. injection</td></tr><tr><td>Carprofen</td><td>Sigma-Aldrich</td><td>SML1713</td><td>5-10mg/kg, for s.c. injection</td></tr><tr><td>4% Paraformaldehyde (PFA)</td><td>Beijing Solarbio Sci-Tech Co.&amp;nbsp;</td><td>P1110</td><td>stocking solution for tissue fixation</td></tr><tr><td>Phosphate buffer (PBS)</td><td>Invitrogen</td><td>10010023</td><td>pH7.4, 500ml in stocking</td></tr><tr><td>Tissue-Tek O.C.T. compound</td><td>Sakura</td><td>4583</td><td>Formulation of water-soluble glycols and resins</td></tr><tr><td>anti-BrdU antibody</td><td>Abcam</td><td>ab6326</td><td>Dilutions:1/800</td></tr><tr><td>anti-c-fos antibody</td><td>Abcam</td><td>ab209794</td><td>Dilutions:1/500</td></tr><tr><td>Goat Anti-Rabbit IgG (Alexa Fluor 568)</td><td>Thermo Fisher Scientific</td><td>A11036</td><td>Dilutions:1/500</td></tr><tr><td>Donkey Anti-Rat IgG (Alexa Fluor 488)</td><td>Jackson ImmunoResearch</td><td>712-546-150</td><td>Dilutions:1/500</td></tr><tr><td>Antifade mounting medium with DAPI</td><td>Vector Laboratories</td><td>H-1200</td><td>Counterstaining with DAPI</td></tr><tr><td>anti-Notch1 antibody (C-20)</td><td>Santa Cruz Biotech</td><td>sc-6014</td><td>Dilutions:1/50</td></tr><tr><td>Donkey Anti-Goat IgG (Alexa Fluor 488)</td><td>Abcam</td><td>ab150073</td><td>Dilutions:1/1000</td></tr></tbody></table>

an invasive method for the activation of the mouse dentate gyrus by high-frequency stimulation

Electrical high-frequency stimulation (HFS), using implanted electrodes targeting various brain regions, has been proven as an effective treatment for various neurological and psychiatric disorders. HFS in the deep region of the brain, also named deep-brain stimulation (DBS), is becoming increasingly important in clinical trials. Recent progress in the field of high-frequency DBS (HF-DBS) surgery has begun to spread the possibility of utilizing this invasive technique to other situations, such as treatment for major depression disorder (MDD), obsessive-compulsive disorder (OCD), and so on. 
Despite these expanding indications, the underlying mechanisms of the beneficial effects of HF-DBS remain enigmatic. To address this question, one approach is to use implanted electrodes that sparsely activate distributed subpopulations of neurons by HFS. It has been reported that HFS in the anterior nucleus of the thalamus could be used for the treatment of refractory epilepsy in the clinic. The underlying mechanisms might be related to the increased neurogenesis and altered neuronal activity. Therefore, we are interested in exploring the physiological alterations by the detection of neuronal activity as well as neurogenesis in the mouse dentate gyrus (DG) before and after HFS treatment. 
In this manuscript, we describe methodologies for HFS to target the activation of the DG in mice, directly or indirectly and in an acute or chronic manner. In addition, we describe a detailed protocol for the preparation of brain slices for c-fos and Notch1 immunofluorescent staining to monitor the neuronal activity and signaling activation and for bromodeoxyuridine (BrdU) labeling to determine the neurogenesis after the HF-DBS induction. The activation of the neuronal activity and neurogenesis after the HF-DBS treatment provides direct neurobiological evidence and potential therapeutic benefits. Particularly, this methodology can be modified and applied to target other interested brain regions such as the basal ganglia and subthalamic regions for specific brain disorders in the clinic.

Following the HF-DBS stimulation to the hippocampal DG subregion directly or the PP subregion to activate the DG indirectly via inserted electrodes using the stereotactic adjustments, the rodents were anesthetized with pentobarbital and sampled 3 h after the last HF-DBS stimulation for the c-fos and Notch1 immunostaining. For the BrdU staining, 36 h after the last BrdU injection after 1 day or 5 days of HF-DBS stimulation, the rodents were anesthetized with pentobarbital...

Watch this Scientific Journal Video about An Invasive Method for the Activation of the Mouse Dentate Gyrus by High-frequency Stimulation at JoVE.com

An Invasive Method for the Activation of the Mouse Dentate Gyrus by High-frequency Stimulation

This protocol shows how to set up a reliable HFS method in mice. Neurons throughout the hippocampal dentate gyrus are electrically stimulated by HFS directly and indirectly in vivo. Neuronal activity and molecular signaling are examined by c-fos and Notch1 immunofluorescent staining, respectively; neurogenesis is quantified by bromodeoxyuridine labeling assay.

Electrical high-frequency stimulation (HFS), using implanted electrodes targeting various brain regions, has been proven as an effective treatment for ...

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Universidad de Cartagena UDEC

Research

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Neuroscience

7.7K Views.  Beijing Institute of Basic Medical Sciences. This protocol shows how to set up a reliable HFS method in mice. Neurons throughout the hippocampal dentate gyrus are electrically stimulated by HFS directly and indirectly in vivo. Neuronal activity and molecular signaling are examined by c-fos and Notch1 immunofluorescent staining, respectively; neurogenesis is quantified by bromodeoxyuridine labeling assay.

Video: An Invasive Method for the Activation of the Mouse Dentate Gyrus by High-frequency Stimulation

Long-term Potentiation of Perforant Pathway-dentate Gyrus Synapse in Freely Behaving Mice

Studies of long-term potentiation of synaptic efficacy, an activity-dependent synaptic phenomenon having properties that make it attractive as a potential cellular mechanism underlying learning and information storage, have long been used to elucidate the physiology of various neuronal circuits in the hippocampus, amygdala, and other limbic and cortical structures. With this in mind, transgenic mouse models of neurological diseases represent useful platforms to conduct long-term&#160;potentiation (LTP) studies to develop a greater understanding of the role of genes in normal and abnormal synaptic communication in neuronal networks involved in learning, emotion and information processing. This article describes methodologies for reliably inducing LTP in the freely behaving mouse. These methodologies can be used in studies of transgenic and knockout freely&#160;behaving mouse models of neurodegenerative diseases.

Transgenic and knockout mouse models of neurological diseases are useful for studying the role of genes in normal and abnormal neurophysiology. This article describes methodologies which can be used to study long-term potentiation, a cellular mechanism which may underlie learning and memory, in transgenic and knockout freely&#160;behaving mouse models of neuropathology.

Studies of long-term potentiation of synaptic efficacy, an activity-dependent synaptic phenomenon having properties that make it attractive as a potential ...

Behavior

Flat-floored Air-lifted Platform: A New Method for Combining Behavior with Microscopy or Electrophysiology on Awake Freely Moving Rodents

It is widely acknowledged that the use of general anesthetics can undermine the relevance of electrophysiological or microscopical data obtained from a living animal&#8217;s brain. Moreover, the lengthy recovery from anesthesia limits the frequency of repeated recording/imaging episodes in longitudinal studies. Hence, new methods that would allow stable recordings from non-anesthetized behaving mice are expected to advance the fields of cellular and cognitive neurosciences. Existing solutions range from mere physical restraint to more sophisticated approaches, such as linear and spherical treadmills used in combination with computer-generated virtual reality. Here, a novel method is described where a head-fixed mouse can move around an air-lifted mobile homecage and explore its environment under stress-free conditions. This method allows researchers to perform behavioral tests (e.g., learning, habituation or novel object recognition) simultaneously with two-photon microscopic imaging and/or patch-clamp recordings, all combined in a single experiment. This video-article describes the use of the awake animal head fixation device (mobile homecage), demonstrates the procedures of animal habituation, and exemplifies a number of possible applications of the method.

This method creates a tangible, familiar environment for the mouse to navigate and explore during microscopic imaging or single-cell electrophysiological recordings, which require firm fixation of the animal&#8217;s head.

It is widely acknowledged that the use of general anesthetics can undermine the relevance of electrophysiological or microscopical data obtained from a ...

Optogenetic Stimulation of the Auditory Nerve

Direct electrical stimulation of spiral ganglion neurons (SGNs) by cochlear implants (CIs) enables open speech comprehension in the majority of implanted deaf subjects1-6. Nonetheless, sound coding with current CIs has poor frequency and intensity resolution due to broad current spread from each electrode contact activating a large number of SGNs along the tonotopic axis of the cochlea7-9. Optical stimulation is proposed as an alternative to electrical stimulation that promises spatially more confined activation of SGNs and, hence, higher frequency resolution of coding. In recent years, direct infrared illumination of&#160;the cochlea has been used to evoke responses in the auditory nerve10. Nevertheless it requires higher energies than electrical stimulation10,11 and uncertainty remains as to the underlying mechanism12. Here we describe a method based on optogenetics to stimulate SGNs with low intensity blue light, using transgenic mice with neuronal expression of channelrhodopsin 2 (ChR2)13 or virus-mediated expression of the ChR2-variant CatCh14. We used micro-light emitting diodes (&#181;LEDs) and fiber-coupled lasers to stimulate ChR2-expressing SGNs through a small artificial opening (cochleostomy) or the round window. We assayed the responses by scalp recordings of light-evoked potentials (optogenetic auditory brainstem response: oABR) or by microelectrode recordings from the auditory pathway and compared them with acoustic and electrical stimulation.

Cochlear implants (CIs) enable hearing by direct electrical stimulation of the auditory nerve. However, poor frequency and intensity resolution limits the quality of hearing with CIs. Here we describe optogenetic stimulation of the auditory nerve in mice as an alternative strategy for auditory research and developing future CIs.

Direct electrical stimulation of spiral ganglion neurons (SGNs) by cochlear implants (CIs) enables open speech comprehension in the majority of implanted ...

Optogenetic Entrainment of Hippocampal Theta Oscillations in Behaving Mice

Extensive data on relationships of neural network oscillations to behavior and organization of neuronal discharge across brain regions call for new tools to selectively manipulate brain rhythms. Here we describe an approach combining projection-specific optogenetics with extracellular electrophysiology for high-fidelity control of hippocampal theta oscillations (5-10 Hz) in behaving mice. The specificity of the optogenetic entrainment is achieved by targeting channelrhodopsin-2 (ChR2) to the GABAergic population of medial septal cells, crucially involved in the generation of hippocampal theta oscillations, and a local synchronized activation of a subset of inhibitory septal afferents in the hippocampus. The efficacy of the optogenetic rhythm control is verified by a simultaneous monitoring of the local field potential (LFP) across lamina of the CA1 area and/or of neuronal discharge. Using this readily implementable preparation we show efficacy of various optogenetic stimulation protocols for induction of theta oscillations and for the manipulation of their frequency and regularity. Finally, a combination of the theta rhythm control with projection-specific inhibition addresses the readout of particular aspects of the hippocampal synchronization by efferent regions.

We describe the use of optogenetics and electrophysiological recordings for selective manipulations of hippocampal theta oscillations (5-10 Hz) in behaving mice. The efficacy of the rhythm entrainment is monitored using local field potential. A combination of opto- and pharmacogenetic inhibition addresses the efferent readout of hippocampal synchronization.

Extensive data on relationships of neural network oscillations to behavior and organization of neuronal discharge across brain regions call for new tools ...

Transcranial Direct Current Stimulation (tDCS) in Mice

Transcranial direct current stimulation (tDCS) is a non-invasive neuromodulation technique proposed as an alternative or complementary treatment for several neuropsychiatric diseases. The biological effects of tDCS are not fully understood, which is in part explained due to the difficulty in obtaining human brain tissue. This protocol describes a tDCS mouse model that uses a chronically implanted electrode allowing the study of the long-lasting biological effects of tDCS. In this experimental model, tDCS changes the cortical gene expression and offers a prominent contribution to the understanding of the rationale for its therapeutic use.

Transcranial Direct Current Stimulation tDCS in Mice

Transcranial direct current stimulation (tDCS) is a therapeutic technique proposed to treat psychiatric diseases. An animal model is essential for understanding the specific biological alterations evoked by tDCS. This protocol describes a tDCS mouse model that uses a chronically implanted electrode.

Transcranial direct current stimulation (tDCS) is a non-invasive neuromodulation technique proposed as an alternative or complementary treatment for ...

Non-invasive Strategies for Chronic Manipulation of DREADD-controlled Neuronal Activity

Chemogenetic strategies have emerged as reliable tools for remote control of neuronal activity. Among these, designer receptors exclusively activated by designer drugs (DREADDs) have become the most popular chemogenetic approach used in modern neuroscience. Most studies deliver the ligand clozapine-N-oxide (CNO) using a single intraperitoneal injection, which is suitable for the acute activation/inhibition of the targeted neuronal population. There are, however, only a few examples of strategies for chronic modulation of DREADD-controlled neurons, the majority of which rely on the use of delivery systems that require surgical intervention. Here, we expand on two non-invasive strategies for delivering the ligand CNO to chronically manipulate neural population in mice. CNO was administered either by using repetitive (daily) eye-drops, or chronically through the animal&#39;s drinking water. These non-invasive paradigms result in robust activation of the designer receptors that persisted throughout the CNO treatments. The methods described here offer alternatives for the chronic DREADD-mediated control of neuronal activity and may be useful for experiments designed to evaluate behavior in freely moving animals, focusing on less-invasive CNO delivery methods.

Here we describe two non-invasive methods to chronically control neuronal activity using chemogenetics in mice. Eye-drops were used to deliver clozapine-N-oxide (CNO) daily. We also describe two methods for prolonged administration of CNO in drinking water. These strategies for chronic neuronal control require minimal intervention reducing animals&#8217; stress.

Chemogenetic strategies have emerged as reliable tools for remote control of neuronal activity. Among these, designer receptors exclusively activated by ...

Stimulating Mouse Brain with a Low-Cost EEG and Millimeter-Sized Coil

Low-Cost Electroencephalographic Recording System Combined with a Millimeter-Sized Coil to Transcranially Stimulate the Mouse Brain In Vivo

A low-cost electroencephalographic (EEG) recording system is proposed here to drive transcranial magnetic stimulation (TMS) of the mouse brain in vivo, utilizing a millimeter-sized coil. Using conventional screw electrodes combined with a custom-made, flexible, multielectrode array substrate, multi-site recording can be carried out from the mouse brain. In addition, we explain how a millimeter-sized coil is produced using low-cost equipment usually found in laboratories. Practical procedures for fabricating the flexible multielectrode array substrate and the surgical implantation technique for screw electrodes are also presented, which are necessary to produce low-noise EEG signals. Although the methodology is useful for recording from the brain of any small animal, the present report focuses on electrode implementation in an anesthetized mouse skull. Furthermore, this method can be easily extended to an awake small animal that is connected with tethered cables via a common adapter and fixed with a TMS device to the head during recording.The present version of the EEG-TMS system, which can include a maximum of 32 EEG channels (a device with 16 channels is presented as an example with fewer channels) and one TMS channel device, is described. Additionally, typical results obtained by the application of the EEG-TMS system to anesthetized mice are briefly reported.

Author Spotlight: Low-Cost Electroencephalographic Recording System Combined with a Millimeter-Sized Coil to Transcranially Stimulate the Mouse Brain In Vivo  

A low-cost electroencephalographic recording system combined with a millimeter-sized coil is proposed to drive transcranial magnetic stimulation of the mouse brain in vivo. Using conventional screw electrodes with a custom-made, flexible, multielectrode array substrate, multi-site recording can be carried out from the mouse brain in response to transcranial magnetic stimulation.

A low-cost electroencephalographic (EEG) recording system is proposed here to drive transcranial magnetic stimulation (TMS) of the mouse brain in vivo, ...

Selective Manipulation of Theta Oscillations in a Mouse Model

Source: Bender, F. et al., <a href="https://appjove.unicartagenaproxy.elogim.com/v/57349">Optogenetic Entrainment of Hippocampal Theta Oscillations in Behaving Mice.</a> J. Vis. Exp (2018)
This video demonstrates a protocol to modulate theta oscillations in a genetically engineered mouse model with light-sensitive ion channels in its GABAergic medial septum neurons. By delivering light pulses to the hippocampus, the protocol aims to synchronize neuronal activity and generate theta oscillations, which are recorded as local field potentials.

Selektywna manipulacja oscylacjami theta w modelu mysim

All procedures involving animal models have been reviewed by the local institutional animal care committee and the JoVE veterinary review board.
PV-Cre ...

JoVE Encyclopedia of Experiments

Neurophysiology

Stimulation and Modulation Techniques

Investigating Auditory Neuronal Responses in an Awake Mouse in the Presence of an Inhibitory Neurotransmitter Blocker

Source: Ayala, Y. A., et al. <a href="https://appjove.unicartagenaproxy.elogim.com/v/53914">Extracellular Recording of Neuronal Activity Combined with Microiontophoretic Application of Neuroactive Substances in Awake Mice.</a> J. Vis. Exp. (2016).
This video demonstrates the study of neuronal activity in a specific region of the mouse brain that processes auditory stimuli. The technique involves applying an inhibitory neurotransmission blocker and comparing neuronal responses to auditory stimuli before and after the drug is applied to the localized brain region.

Badanie słuchowych reakcji neuronalnych u obudzonej myszy w obecności hamującego blokera neuroprzekaźników

All procedures involving animal models have been reviewed by the local institutional animal care committee and the JoVE veterinary review board.
1. Drugs ...

Implanting an Electrode Array and Optic Fiber into the Mouse Hippocampus

Source: Bender, F., et. al., <a href="https://appjove.unicartagenaproxy.elogim.com/v/57349">Optogenetic Entrainment of Hippocampal Theta Oscillations in Behaving Mice.</a> J. Vis. Exp. (2018)
This video demonstrates the stereotactic implantation of a wire electrode array and an optic fiber in the hippocampus of a mouse model.

Wszczepianie układu elektrod i światłowodu do hipokampa myszy

All procedures involving animal models have been reviewed by the local institutional animal care committee and the JoVE veterinary review board.
1. ...