This work was supported by the intramural research program at the National Institute of Mental Health (ZIA MH002964-02). We would like to thank the support of the NIMH IRP Rodent Behavioral Core (ZIC MH002952).

All animals were handled in accordance with guidelines of the Animal Care and Use Committees of the National Institute of Mental Health (NIMH). All efforts were made to minimize the pain and the number of animals used.
1. Adeno-associated virus injections in the hippocampus
NOTE: Wild type male mice of mixed background (B6/129 F1 hybrid, 3 months old) were for stereotaxically injected with&#160;an AAV encoding the M3 muscarinic receptor (hM3Dq) into the hippocampus. During the entire experiment, mice were single-housed, under a regular 12 h light: 12 h dark (T24) cycle, with access to food and water ad libitum.
<ol>
	<li>Before performing stereotaxic surgeries, clean and sterilize the stereotaxic frame and all needed instruments. 
	NOTE: Surgical drapes could be used to maintain a sterile field and reduce mouse&#39;s heat loss.</li>
	<li>Deeply anesthetize the mouse using isoflurane. To do this, first adjust the oxygen flow meter to approximately 1.5 L/ min, and then adjust the isoflurane vaporizer to approximately 3-5% for induction and approximately 1-3% for maintenance.
	<ol>
		<li>To ensure that the animal is fully unconscious, pinch the mouse&#8217;s paw; the animal is properly anesthetized when the flinching response to pinch is absent.</li>
	</ol>
	</li>
	<li>Place the mouse on a heating pad to maintain the stability of the mouse&#8217;s body temperature.</li>
	<li>Shave the top of the head and fix the head of the mouse to the stereotaxic frame. Then, apply ocular protective lubricant on the eyes, clean the surface by scrubbing with povidone-iodine and 70% ethanol, and expose the skull using a sterile scalpel.</li>
	<li>Calibrate the frame to bregma point, then drill at a medial-lateral coordinate of 2.9 mm and an anterior-posterior coordinate -2.7 mm to target the hippocampus. 
	NOTE: If other brain target needs to be injected, determine the desired coordinates for injection using the Paxinos and Franklin mouse atlas16.</li>
	<li>Once the brain is exposed, unilaterally inject 90 nL of the AAV at the dorsal-ventral depth of -3.0 mm in the hippocampus using a microinjector and pulled microcapillary pipettes (Figure 1A). 
	NOTE: See Table of Materials for the titer of AAV used in this experiment. For other brain areas, adjust the AAV volume of injection as needed.</li>
	<li>At the end of the surgical procedure, close the incision with nylon sutures and apply topical antibiotics to the wound site.</li>
	<li>Administer analgesics (buprenorphine, 0.1 mg/kg) systemically immediately following surgery, and 4-6 hours after.</li>
	<li>Beginning 4 weeks post-injection, subject mice to any of the paradigms described in the following section to chronically control neurons expressing the designer excitatory receptor.</li>
</ol>
2. Repetitive CNO delivery using eye-drops
<ol>
	<li>Acclimate the animals to handling by scruffing each mouse 3 min daily for 3-4 days prior to the administration of eye-drops.</li>
	<li>Dissolve Clozapine-N-oxide (CNO, 5 mg) in 1 mL of sterile 0.9% saline solution (stock solution: 5 mg CNO/ mL). Keep the solution refrigerated at 4 &#176;C.</li>
	<li>Weigh each mouse before starting the experiment to determine the amount of CNO to be delivered. Use 1-3 &#956;L drop (per eye) to achieve 1.0 mg CNO/ kg body weight. 
	NOTE: As an example, a 20 g mouse should receive bilateral (2&#160;&#956;L each) eye-drops.</li>
	<li>Deliver the eye-drops during the inactive (light) phase of mice, 2 h before lights turn off (zeitgeber time (ZT) 10). In cases where CNO needs to be delivered during the active (dark) phase of mice, ensure the presence of dim red light for proper animal handling. 
	NOTE: Precautions should be taken to avoid disrupting the circadian (and light/dark) cycle of experimental animals.
	<ol>
		<li>Using a P10 micropipette, load the required amount (1-3 &#956;L) of CNO solution to achieve 1.0 mg CNO/ kg. 
		NOTE: Use a new and sterile pipette tip for each eye-drop. In this set of experiments, bilateral eye-drops of CNO were performed; however, if a lower CNO concentration is required, unilateral eye-drops could be also applied.</li>
		<li>Immobilize the mouse via scruff.</li>
		<li>Slowly expel the solution until a stable droplet forms on the pipette tip.</li>
		<li>Carefully bring the droplet close to the cornea of the mouse&#8217;s eye until the solution is delivered. The pipette tip should never contact the mouse&#8217;s eye.</li>
		<li>Release the mouse, placing it back in its home cage.</li>
	</ol>
	</li>
	<li>Repeat this procedure every day for 5 days. 
	NOTE: This duration can be adjusted as per the experimental requirements.</li>
	<li>For control experiments, use AAV/DREADD-injected mice subjected to sham treatment (eye-drops containing only saline solution), and mice injected with an empty vector (e.g., AAV/mCherry) exposed to the described CNO eye-drops protocol.</li>
</ol>
3. Chronic CNO treatment delivered through drinking water
<ol>
	<li>Make small bottles using 50 mL (plastic) conical tubes and rubber stopper spouts; cover with aluminum foil to avoid any light-mediated effects on CNO stability.</li>
	<li>Three days before starting with the CNO treatment, replace regular water bottles with small bottles, containing 10 mL of regular water, to allow mice to acclimate to them. Secure the bottles to the cages using tape.</li>
	<li>Measure the daily water consumption for each mouse.</li>
	<li>Weigh each mouse before starting the experiment.</li>
	<li>Dissolve Clozapine-N-oxide (CNO, 5 mg) in 1 mL of 0.9% sterile saline solution. Refrigerate the stock solution at 4 &#176;C.</li>
	<li>Use the body weight and the average amount of water consumed to define the concentration of CNO solution to achieve 1.0 mg CNO/ kg (body weight). 
	NOTE: Adult male mice (~20 g body weight) consume ~5 mL of water per day (Figure 2A). Therefore, to achieve a CNO concentration of 1 mg CNO/ kg, 6.4 &#956;L of CNO stock solution should be added to a final volume of 8 mL of water (final concentration: 4 &#956;g CNO/ mL). Thus, the dose of CNO for a 20 g animal that drinks 5 mL water per day results in 1 mg CNO/ kg.</li>
	<li>Determine the optimal CNO dose that displays the maximum effectiveness with minimal CNO concentration by testing a range of concentrations. Perform a dose response analysis to determine the optimal CNO dose for the drinking water method. 
	NOTE: The following CNO doses were tested for this experiment: 1.0 mg/ mL, 0.5 mg/ mL, 0.25 mg/ mL, 0.1 mg/ mL, and saline. 1.0 mg CNO/ kg was first tested, based on i.p. and eye-drops protocols.</li>
	<li>On day 1, fill the bottle with 8 mL of regular water and add the required amount of CNO. 
	NOTE: This amount of water is enough for 24 h of ad libitum water access for an adult male mouse. In case other rodent species are used, first measure the amount of water consumed daily to determine the volume needed.</li>
	<li>Monitor the health of the animals throughout the protocol to ensure that there are no adverse side effects caused by water + CNO consumption.</li>
	<li>After 24 h, replace the bottles with fresh water + CNO solution. Record the volume consumed during the previous day.</li>
	<li>Dispose of the water + CNO solution that was not consumed in waste containers. Discard plastic bottles and replace the rubber stoppers every day, after sanitizing them according to the animal facility guidelines. 
	NOTE: Do not mix aqueous wastes with organic solvents. Contact the Chemical Disposal Service for instructions for storage and pick-up.</li>
	<li>Replace the bottles at the same time every day for 5 days. 
	NOTE: This duration can be adjusted as per the experimental requirements.</li>
	<li>Include control groups, as described in step 2.6.</li>
</ol>
4. Restricted CNO treatment using mice&#8217;s preference for sucrose
<ol>
	<li>3 days before starting with the CNO treatment, place a small bottle containing 10 mL of water + 1% sucrose on the cage, preferably away from the original water bottle. 
	NOTE: Use the same small bottles described in Step 3.1.</li>
	<li>Expose animals to water + 1% sucrose during the last portion of their active phase (ZT 18 &#8211; 24). After this exposure, remove the bottle with water + sucrose from the cage. 
	NOTE: Different time windows of CNO delivery could be used. Additionally, mice could be placed under an inverted light/dark cycle, where the onset of light occurs in the evening hours to facilitate the CNO delivery.</li>
	<li>Measure the daily water + 1% sucrose consumption for each mouse.</li>
	<li>Weigh each mouse before starting the experiment.</li>
	<li>Use the body weight and the average amount of water + 1% sucrose consumed to determine the dose of CNO solution to achieve 1.0 mg CNO/ kg (body weight). 
	NOTE: The optimal CNO dose that displays the maximum effectiveness with minimal CNO concentration should be tested, as explained in step 3.6.</li>
	<li>On day 1, fill bottles with 5 mL of water + 1% sucrose + CNO (1 mg CNO/ Kg) and place them on the cage (always at the same location) during the determined time window.</li>
	<li>At the end of the restricted time window, remove the bottles and measure the amount of water + sucrose + CNO consumed. 
	NOTE: Materials and solutions are sanitized or discarded as previously described in step 3.11.</li>
	<li>Repeat this procedure every day for 5 days. 
	NOTE: This duration can be adjusted as per the experimental requirements.</li>
	<li>Include a control group, as described in step 2.6.</li>
</ol>
5. Data analysis
<ol>
	<li>Perfuse mice intracardially with 4% paraformaldehyde (PFA) either 2 or 6 h after receiving the last repetitive (5th day) CNO eye-drop. When CNO is delivered through drinking water, replace CNO + water with water at the end of the mouse&#8217;s active phase, then perfuse the mouse after either 2 or 6 h post-CNO access. 
	NOTE: If light exposure could affect the c-Fos induction in the area of interest, keep mice in constant darkness during the last day of the experiments, and before the perfusion.</li>
	<li>Carefully dissect the brain out and submerge in 4% PFA solution for 9-12h.</li>
	<li>After PFA fixation, cryoprotect the brain tissue using a 30% sucrose solution (wait until the brain sinks), then section the brain using a cryostat.</li>
	<li>Transfer the coronal brain sections (35 &#956;m) into a solution containing 1x PBS, 10% bovine serum albumin, and 0.3% Triton X-100 for 1 h at room temperature.</li>
	<li>Incubate the brain sections with an anti-c-Fos (1:2500) antibody solution at 4 &#176;C overnight with constant agitation.</li>
	<li>After 3 washes of 5 min each with a solution containing 1x PBS and 0.3% Triton X-100, incubate the samples with an Alexa-conjugated secondary antibody (1:500) solution for 1 h at room temperature away from light and with constant agitation.</li>
	<li>Obtain digital images using a confocal microscope. Assemble and process captured images with a photo editing and analysis software (e.g., Adobe Photoshop).</li>
	<li>For data analysis, outline and measure the AAV-infected area (mCherry(+) cells) using ImageJ software, and quantify the number of c-Fos(+) cells within this region to obtain the number of activated cells per area. Combine the results obtained from 3 separate sections per animal.</li>
</ol>

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The authors have nothing to disclose.

DREADDs have emerged as a popular and effective approach to remotely manipulate neuronal activity17. The design of alternative strategies for CNO delivery will broadly increase the spectrum of options available for specific experimental settings. In addition, non-invasive strategies for the delivery of CNO minimize any potential misinterpretation of results by reducing adverse side effects that can directly impact the animal&#8217;s health. Here, we described two non-invasive strategies for CNO de...

Technical advances in the field of neuroscience have allowed scientists to precisely identify and control the activity of particular neuronal populations1. This has contributed to better understand the basis of neuronal circuits and their impact on animal behavior, as well as, revising established dogmas2,3. Among these novel tools, optogenetic and chemogenetic strategies have had a profound impact not only on the quality of discoveries but also on the way experiments are conceived and designed4. In the present manuscript, we focus on chemogenetic strategies for controlling the activation of neurons via engineered receptor-ligand strategies. Designer receptors exclusively activated by designer drugs (DREADDs) represent one of the most popular chemogenetic tools for the remote control of neuronal activity, as reviewed by Roth 20165. DREADDs utilize modified muscarinic acetylcholine receptors that are specifically activated by an inert ligand, clozapine-N-oxide (CNO)6.
Most studies use CNO administered by intraperitoneal (i.p.) injections, which effectively controls the dosage and timing of engineered receptors activation in an acute fashion. However, when repetitive or chronic DREADD activation is required, the use of multiple i.p. injections become unfeasible. To address this issue, different strategies for the chronic CNO delivery have been reported, including implanted minipumps7 and intracranial cannulas8,9. To different extents, all these strategies cause the animals stress and pain10, and require a surgical intervention that could also have a direct impact on the behavioral responses to be tested11. Here, we describe three non-invasive strategies for the chronic CNO delivery.
For this purpose, mice were stereotaxically injected in the hippocampus with an adeno-associated virus (AAV) encoding an engineered version of the excitatory M3 muscarinic receptor (hM3Dq) that when activated by the ligand CNO leads to the burst-like firing of neurons6. It was previously shown that a single eye-drop containing CNO can effectively elicit a robust activation of DREADD-expressing neurons12. Here we describe a modified method for the repetitive delivery of eye drops. To achieve chronic and sustained control of the designer receptors, we next describe a non-invasive strategy to deliver CNO to mice through the drinking water. Finally, we describe an alternative paradigm for delivering CNO in drinking water during a restricted time window. Mice locomotor activity, as well as drinking behavior and the consumption of sweet caloric solutions, are mostly restricted to the dark portion of the light/dark cycle13,14. Therefore, we adopted a protocol based on the mouse&#8217;s preference for sucrose. By measuring the induction of the immediate-early gene c-Fos in AAV-infected cells, as a readout for neuronal activation12,15, we found that these CNO delivery strategies robustly activate DREADD-controlled neurons over extended durations.

<table>
	<tbody>
		<tr>
			<td>BSA</td>
			<td>Sigma life science</td>
			<td>#A2153-100G</td>
			<td>Lyophilized powder &#8805;96% (agarose gel electrophoresis)</td>
		</tr>
		<tr>
			<td>C57BL/6J mice</td>
			<td>The Jackson laboratory</td>
			<td>#000664</td>
			<td>male mice, 3 months old</td>
		</tr>
		<tr>
			<td>Capillaries</td>
			<td>Drummond Scientific Company</td>
			<td>#3-000-203-G/X</td>
			<td>Outer diameter: 1.14 in.</td>
		</tr>
		<tr>
			<td>Clozapine-N-oxide</td>
			<td>Sigma</td>
			<td>#C0832</td>
			<td>5mg</td>
		</tr>
		<tr>
			<td>Forane</td>
			<td>Baxter</td>
			<td>#NDC 10019-360-60</td>
			<td>Isoflurane, USP</td>
		</tr>
		<tr>
			<td>Microinjector III</td>
			<td>Drummond Scientific Company</td>
			<td>#3-000-207</td>
			<td>Nanoject III - Programmable Nanoliter Injector</td>
		</tr>
		<tr>
			<td>Mounting media</td>
			<td>Invitrogen</td>
			<td>#P36930</td>
			<td>Prolong Gold antifade reagent</td>
		</tr>
		<tr>
			<td>Paraformaldehyde</td>
			<td>Electron Microscopy Sciences</td>
			<td>#15710</td>
			<td>16% aqueous solution (methanol free), 10 ml</td>
		</tr>
		<tr>
			<td>Primary c-Fos Antibody</td>
			<td>Cell signaling technology</td>
			<td>#2250S</td>
			<td>c-Fos (9F6) Rabbit mAb (100&#181;l)</td>
		</tr>
		<tr>
			<td>rAAV5/hSyn-hm3D-mCherry</td>
			<td>UNC Vector Core</td>
			<td></td>
			<td>Titer: ~3x10e12 vg/mL</td>
		</tr>
		<tr>
			<td>rAAV5/hSyn-mCherry</td>
			<td>UNC Vector Core</td>
			<td></td>
			<td>Titer: ~3x10e12 vg/mL</td>
		</tr>
		<tr>
			<td>Secondary Antibody</td>
			<td>Invitrogen</td>
			<td>#A21206</td>
			<td>Alexa Fluor TM 488 Donkey anti-rabbit IgG(H+L), 2mg/ml</td>
		</tr>
		<tr>
			<td>Triton X-100</td>
			<td>americanbio.com</td>
			<td>#AB02025-00100</td>
			<td></td>
		</tr>
	</tbody>
</table>

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.

We observed that repetitive CNO delivery using eye-drops elicited a robust induction of c-Fos expression in most infected neurons (Figure 1C), showing that the effectiveness of CNO delivery is sustained during the repetitive exposure. Furthermore, a significant induction of c-Fos was observed in samples collected 2 h after CNO treatment, compared to samples obtained 6 h after CNO exposure (Figures 1D-E), demonstrating that changes induced by CNO are time-dependent.
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Non-invasive Strategies for Chronic Manipulation of DREADD-controlled Neuronal Activity

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 ...

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Neuroscience

13.3K Views.  National Institute of Mental Health. 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’ stress.

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

Surgical Implantation of Chronic Neural Electrodes for Recording Single Unit Activity and Electrocorticographic Signals

The success of long-term electrophysiological recordings often depends on the quality of the implantation surgery. Here we provide useful information for surgeons who are learning the process of implanting electrode systems. We demonstrate the implantation procedure of both a penetrating and a surface electrode. The surgical process is described from start to finish, including detailed descriptions of each step throughout the procedure. It should also be noted that this video guide is focused towards procedures conducted in rodent models and other small animal models. Modifications of the described procedures are feasible for other animal models.

We provide useful information for surgeons who are learning the process of implanting chronic neural recording electrodes. Techniques for both penetrating and surface electrode systems are described in a rodent animal model.

The success of long-term electrophysiological recordings often depends on the quality of the implantation surgery.  Here we provide useful information for ...

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

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

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

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

A Procedure for Implanting Organized Arrays of Microwires for Single-unit Recordings in Awake, Behaving Animals

In vivo electrophysiological recordings in the awake, behaving animal provide a powerful method for understanding neural signaling at the single-cell level. The technique allows experimenters to examine temporally and regionally specific firing patterns in order to correlate recorded action potentials with ongoing behavior. Moreover, single-unit recordings can be combined with a plethora of other techniques in order to produce comprehensive explanations of neural function. In this article, we describe the anesthesia and preparation for microwire implantation. Subsequently, we enumerate the necessary equipment and surgical steps to accurately insert a microwire array into a target structure. Lastly, we briefly describe the equipment used to record from each individual electrode in the array. The fixed microwire arrays described are well-suited for chronic implantation and allow for longitudinal recordings of neural data in almost any behavioral preparation. We discuss tracing electrode tracks to triangulate microwire positions as well as ways to combine microwire implantation with immunohistochemical techniques in order to increase the anatomical specificity of recorded results.

Implanting organized arrays of microwires for use in single-unit electrophysiological recordings presents a number of technical challenges.&#160;Methods for performing this technique and the equipment necessary are described.&#160;Also, the beneficial use of organized microwire arrays to record from distinct neural subregions with high spatial selectivity is discussed.

In vivo electrophysiological recordings in the awake, behaving animal provide a powerful method for understanding neural signaling at the single-cell ...

Genetic Manipulation of Cerebellar Granule Neurons In Vitro and In Vivo to Study Neuronal Morphology and Migration

Developmental events in the brain including neuronal morphogenesis and migration are highly orchestrated processes. In vitro and in vivo analyses allow for an in-depth characterization to identify pathways involved in these events. Cerebellar granule neurons (CGNs) that are derived from the developing cerebellum are an ideal model system that allows for morphological analyses. Here, we describe a method of how to genetically manipulate CGNs and how to study axono- and dendritogenesis of individual neurons. With this method the effects of RNA interference, overexpression or small molecules can be compared to control neurons. In addition, the rodent cerebellar cortex is an easily accessible in vivo system owing to its predominant postnatal development. We also present an in vivo electroporation technique to genetically manipulate the developing cerebella and describe subsequent cerebellar analyses to assess neuronal morphology and migration.

Genetic Manipulation of Cerebellar Granule Neurons In Vitro and In Vivo to Study Neuronal Morphology and Migration

Neuronal morphogenesis and migration are crucial events underlying proper brain development. Here, we describe methods to genetically manipulate cultured cerebellar granule neurons and the developing cerebellum for the assessment of morphology and migratory characteristics of neurons.

Developmental events in the brain including neuronal morphogenesis and migration are highly orchestrated processes. In vitro and in vivo analyses allow ...

Inducing Plasticity of Astrocytic Receptors by Manipulation of Neuronal Firing Rates

Close to two decades of research has established that astrocytes in situ and in vivo express numerous G protein-coupled receptors (GPCRs) that can be stimulated by neuronally-released transmitter. However, the ability of astrocytic receptors to exhibit plasticity in response to changes in neuronal activity has received little attention. Here we describe a model system that can be used to globally scale up or down astrocytic group I metabotropic glutamate receptors (mGluRs) in acute brain slices. Included are methods on how to prepare parasagittal hippocampal slices, construct chambers suitable for long-term slice incubation, bidirectionally manipulate neuronal action potential frequency, load astrocytes and astrocyte processes with fluorescent Ca2+ indicator, and measure changes in astrocytic Gq GPCR activity by recording spontaneous and evoked astrocyte Ca2+ events using confocal microscopy. In essence, a &#8220;calcium roadmap&#8221; is provided for how to measure plasticity of astrocytic Gq GPCRs. Applications of the technique for study of astrocytes are discussed. Having an understanding of how astrocytic receptor signaling is affected by changes in neuronal activity has important implications for both normal synaptic function as well as processes underlying neurological disorders and neurodegenerative disease.

Here we describe an adaptation of protocols used to induce homeostatic plasticity in neurons for the study of plasticity of astrocytic G protein-coupled receptors. Recently used to examine changes in astrocytic group I mGluRs in juvenile mice, the method can be applied to measure scaling of various astrocytic GPCRs, in tissue from adult mice in situ and in vivo, and to gain a better appreciation of the sensitivity of astrocytic receptors to changes in neuronal activity.

Close to two decades of research has established that astrocytes in situ and in vivo express numerous G protein-coupled receptors (GPCRs) that can be ...

Non-invasive Parenchymal, Vascular and Metabolic High-frequency Ultrasound and Photoacoustic Rat Deep Brain Imaging

Photoacoustics and high frequency ultrasound stands out as powerful tools for neurobiological applications enabling high-resolution imaging on the central nervous system of small animals. However, transdermal and transcranial neuroimaging is frequently affected by low sensitivity, image aberrations and loss of space resolution, requiring scalp or even skull removal before imaging. To overcome this challenge, a new protocol is presented to gain significant insights in brain hemodynamics by photoacoustic and high-frequency ultrasounds imaging with the animal skin and skull intact. The procedure relies on the passage of ultrasound (US) waves and laser directly through the fissures that are naturally present on the animal cranium. By juxtaposing the imaging transducer device exactly in correspondence to these selected areas where the skull has a reduced thickness or is totally absent, one can acquire high quality deep images and explore internal brain regions that are usually difficult to anatomically or functionally describe without an invasive approach. By applying this experimental procedure, significant data can be collected in both sonic and optoacoustic modalities, enabling to image the parenchymal and the vascular anatomy far below the head surface. Deep brain features such as parenchymal convolutions and fissures separating the lobes were clearly visible. Moreover, the configuration of large and small blood vessels was imaged at several millimeters of depth, and precise information were collected about blood fluxes, vascular stream velocities and the hemoglobin chemical state. This repertoire of data could be crucial in several research contests, ranging from brain vascular disease studies to experimental techniques involving the systemic administration of exogenous chemicals or other objects endowed with imaging contrast enhancement properties. In conclusion, thanks to the presented protocol, the US and PA techniques become an attractive noninvasive performance-competitive means for cortical and internal brain imaging, retaining a significant potential in many neurologic fields.

The present work describes a new protocol to perform non-invasive high-frequency ultrasound and photoacoustic based imaging on rat brain, to efficiently visualize deep subcortical regions and their vascular patterns by directing signals on skull foramina naturally present on animal cranium.

Photoacoustics and high frequency ultrasound stands out as powerful tools for neurobiological applications enabling high-resolution imaging on the central ...

Non-invasive Assessment of Changes in Corticomotoneuronal Transmission in Humans

The corticospinal pathway is the major pathway connecting the brain with the muscles and is therefore highly relevant for movement control and motor learning. There exists a number of noninvasive electrophysiological methods investigating the excitability and plasticity of this pathway. However, most methods are based on quantification of compound potentials and neglect that the corticospinal pathway consists of many different connections that are more or less direct. Here, we present a method that allows testing excitability of different fractions of the corticospinal transmission. This so called H-reflex conditioning technique allows one to assess excitability of the fastest (monosynaptic) and also polysynaptic corticospinal pathways. Furthermore, by using two different stimulation sites, the motor cortex and the cervicomedullary junction, it allows not only differentiation between cortical and spinal effects but also assessment of transmission at the corticomotoneural synapse. In this manuscript, we describe how this method can be used to assess corticomotoneural transmission after low-frequency repetitive transcranial magnetic stimulation, a method that was previously shown to reduce excitability of cortical cells. Here we demonstrate that not only cortical cells are affected by this repetitive stimulation but also transmission at the corticomotoneuronal synapse at the spinal level. This finding is important for the understanding of basic mechanisms and sites of neuroplasticity. Besides investigation of basic mechanisms, the H-reflex conditioning technique may be applied to test changes in corticospinal transmission following behavioral (e.g., training) or therapeutic interventions, pathology or aging and therefore allows a better understanding of neural processes that underlie movement control and motor learning.

The aim of the present study was to assess changes in transmission at the corticomotoneuronal synapses in humans after repetitive transcranial magnetic stimulation. For this purpose, an electrophysiological method is introduced that allows assessment of pathway specific corticospinal transmission, i.e. differentiation of fast, direct corticospinal pathways from polysynaptic connections.

The corticospinal pathway is the major pathway connecting the brain with the muscles and is therefore highly relevant for movement control and motor ...

Recording Temperature-induced Neuronal Activity through Monitoring Calcium Changes in the Olfactory Bulb of Xenopus laevis

The olfactory system, specialized in the detection, integration and processing of chemical molecules is likely the most thoroughly studied sensory system. However, there is piling evidence that olfaction is not solely limited to chemical sensitivity, but also includes temperature sensitivity. Premetamorphic Xenopus laevis are translucent animals, with protruding nasal cavities deprived of the cribriform plate separating the nose and the olfactory bulb. These characteristics make them well suited for studying olfaction, and particularly thermosensitivity. The present article describes the complete procedure for measuring temperature responses in the olfactory bulb of X. laevis larvae. Firstly, the electroporation of olfactory receptor neurons (ORNs) is performed with spectrally distinct dyes loaded into the nasal cavities in order to stain their axon terminals in the bulbar neuropil. The differential staining between left and right receptor neurons serves to identify the &#947;-glomerulus as the only structure innervated by contralateral presynaptic afferents. Secondly, the electroporation is combined with focal bolus loading in the olfactory bulb in order to stain mitral cells and their dendrites. The 3D brain volume is then scanned under line-illumination microscopy for the acquisition of fast calcium imaging data while small temperature drops are induced at the olfactory epithelium. Lastly, the post-acquisition analysis allows the morphological reconstruction of the thermosensitive network comprising the &#947;-glomerulus and its innervating mitral cells, based on specific temperature-induced Ca2+ traces. Using chemical odorants as stimuli in addition to temperature jumps enables the comparison between thermosensitive and chemosensitive networks in the olfactory bulb.

Recording Temperature-induced Neuronal Activity through Monitoring Calcium Changes in the Olfactory Bulb of Xenopus laevis

Here we describe a protocol for measuring and analyzing temperature responses in the olfactory bulb of Xenopus laevis. Olfactory receptor neurons and mitral cells are differentially stained, after which calcium changes are recorded, reflecting a sensitivity of some neural networks in the bulb to temperature drops induced at the nose.

The olfactory system, specialized in the detection, integration and processing of chemical molecules is likely the most thoroughly studied sensory system. ...

Chronic Transcranial Electrical Stimulation and Intracortical Recording in Rats

Transcranial electrical stimulation (TES) is a powerful and relatively simple approach to diffusely influence brain activity either randomly or in a closed-loop event-triggered manner. Although many studies are focusing on the possible benefits and side-effects of TES in healthy and pathologic brains, there are still many fundamental open questions regarding the mechanism of action of the stimulation. Therefore, there is a clear need for a robust and reproducible method to test the acute and the chronic effects of TES in rodents. TES can be combined with regular behavioral, electrophysiological, and imaging techniques to investigate neuronal networks in vivo. The implantation of transcranial stimulation electrodes does not impose extra constraints on the experimental design while it offers a versatile, flexible tool to manipulate brain activity. Here we provide a detailed, step-by-step protocol to fabricate and implant transcranial stimulation electrodes to influence brain activity in a temporally constrained manner for months.

This detailed protocol describes transcranial stimulation electrode placement on the temporal bone in order to investigate the short- and long-term effects of transcranial electrical stimulation in freely moving rats.

Transcranial electrical stimulation (TES) is a powerful and relatively simple approach to diffusely influence brain activity either randomly or in a ...

Non-Invasive Modulation and Robotic Mapping of Motor Cortex in the Developing Brain

Mapping the motor cortex with transcranial magnetic stimulation (TMS) has potential to interrogate motor cortex physiology and plasticity but carries unique challenges in children. Similarly, transcranial direct current stimulation (tDCS) can improve motor learning in adults but has only recently been applied to children. The use of tDCS and emerging techniques like high&#8211;definition tDCS (HD-tDCS) require special methodological considerations in the developing brain. Robotic TMS motor mapping may confer unique advantages for mapping, particularly in the developing brain. Here, we aim to provide a practical, standardized approach for two integrated methods capable of simultaneously exploring motor cortex modulation and motor maps in children. First, we describe a protocol for robotic TMS motor mapping. Individualized, MRI-navigated 12x12 grids centered on the motor cortex guide a robot to administer single-pulse TMS. Mean motor evoked potential (MEP) amplitudes per grid point are used to generate 3D motor maps of individual hand muscles with outcomes including map area, volume, and center of gravity. Tools to measure safety and tolerability of both methods are also included. Second, we describe the application of both tDCS and HD-tDCS to modulate the motor cortex and motor learning. An experimental training paradigm and sample results are described. These methods will advance the application of non-invasive brain stimulation in children.

We demonstrate protocols for the modulation (tDCS, HD-tDCS) and mapping (robotic TMS) of the motor cortex in children.

Mapping the motor cortex with transcranial magnetic stimulation (TMS) has potential to interrogate motor cortex physiology and plasticity but carries ...