The project was funded by&#160;a Research Initiative Award from the American Epilepsy Society, and an award from the Arizona Biomedical Research Commission to S.L.M., as well as a challenge grant from Research to Prevent Blindness Inc. to the Department of Ophthalmology at SUNY Downstate Health Sciences University, the New York State Empire Innovation Program, and further grants from the National Science Foundation (0726113, 0852636, &#38; 1523614), the Barrow Neurological Foundation, Mrs. Marian Rochelle, Mrs. Grace Welton, and Dignity Health SEED awards, and by federal grants from the National Science Foundation (0726113, 0852636, &#38; 1523614) and by the National Institute of Health (Awards R01EY031971 AND R01CA258021), to S.L.M and S.M.C.&#160;<span lang="EN-US" style="font-size: 12pt; line-height: 107%; font-family: Calibri, sans-serif; color: rgb(34, 34, 34); background-image: initial; background-position: initial; background-size: initial; background-repeat: initial; background-attachment: initial; background-origin: initial; background-clip: initial;">This work was also supported by the Office of the Assistant Secretary of Defense for Health Affairs under Award No. W81XWH-15-1-0138, to S.L.M..<span lang="EN-US" style="font-size: 12pt; line-height: 107%; font-family: Calibri, sans-serif; color: red; background-image: initial; background-position: initial; background-size: initial; background-repeat: initial; background-attachment: initial; background-origin: initial; background-clip: initial;">&#160;<span lang="EN-US" style="font-size: 12pt; line-height: 107%; font-family: Calibri, sans-serif; color: rgb(34, 34, 34); background-image: initial; background-position: initial; background-size: initial; background-repeat: initial; background-attachment: initial; background-origin: initial; background-clip: initial;">L.-C. was supported by a Jos&#233; Castillejo fellowship from the Spanish Ministry of Education. We thank O. Caballero and M. Ledo for their technical advice and assistance.&#160;

The protocol follows the NIH Guidelines for the Care and Use of Laboratory Animals. All procedures were approved by the Barrow Neurological Institute&#8217;s Institutional Animal Care and Use Committee.
1. Stereotaxic positioning for craniotomy
<ol>
	<li>Weigh and then anesthetize the mouse with a ketamine-xylazine (100 mg/kg&#8211;10 mg/kg i.p.) cocktail. Ensure that the animal is fully anesthetized by observing its lack of reaction to tail and/or toe pinch.</li>
	<li>Place a heating pad under the mouse and use a rectal thermometer to continuously monitor its body temperature during the initial anesthetized implantation surgery (Figure 2A).</li>
	<li>Apply ophthalmic ointment to prevent eye dryness.</li>
	<li>Stabilize the head of the animal in a rodent stereotaxic frame, outfitted with a mouse adapter. To properly orient the mouse in the stereotaxic frame, place the animal&#8217;s teeth inside the hole of the bite-bar (Figure 2A and Figure 3A, item g). Tighten the nose-clamp until it is snug (Figure 2A,B and Figure 3A, item h). The mouse&#8217;s body positioning should be as straight as possible (as in Figure 2A).
	<ol>
		<li>Further secure the animal&#8217;s head using mouse ear-bars with jaw holder cuffs (Figure 2A and Figure 3A, item i), to firmly clamp the zygomatic processes of the skull. 
		NOTE: Once secured, the mouse&#8217;s head should have no range of horizontal motion.</li>
	</ol>
	</li>
	<li>Once the mouse is adjusted to the stereotaxic frame (Figure 3B, item j), clean and sterilize the scalp by wiping it with chlor-hexedine scrub 2-3 times, each time followed by an alcohol rinse. Then, apply topical lidocaine on top of the head.</li>
	<li>Make an incision along the midline from posterior to anterior (start incision at the base of skull and complete it between the eyes; 12-15 mm) along the sagittal midline of the skull, using either fine scissors or a blade. Retract the borders of the scalp with four 28 mm bulldog serrefine clamps to expose the skull and maximize the working area (Figure 2B and Figure 3C).</li>
	<li>Use a scalpel or microspatula (Figure 3C) to scrape the periosteum to the incision&#8217;s edges. Use tissue-separating scissor or forceps (Figure 3C) to carefully retract the muscles at the back of the neck (Figure 2B).</li>
	<li>Dry the top of the skull with a cotton-tipped applicator, moistened with alcohol or 30% hydrogen peroxide, to optimize visualization of the cranial sutures. Apply saline to the skull once the connective tissue has been removed.</li>
	<li>Once the mouse&#8217;s head is stabilized in the stereotaxic apparatus, place a syringe needle tip (21 G) in the electrode holder (Figure 3B, item l), and attach the electrode holder to the stereotaxic micromanipulator, lowering the needle tip to the level of the skull (Figure 2D).</li>
	<li>Adjust the medial-lateral direction of the skull by measuring the height of the skull at any distance posterior to bregma (i.e., -2.0 mm), and two equal distances lateral from the midline on either side of the skull (i.e., &#177; 1.5 mm) (Figure 2D). Aim for a height difference across locations that is less than 0.05 mm in the dorsal-ventral direction.
	<ol>
		<li>If greater than 0.05 mm, rotate the mouse&#8217;s skull using the bite bar, or reposition the mouse&#8217;s head by loosening the ear-bars carefully, laterally shifting the head and ear-bars as needed, and then retightening the ear-bars.</li>
	</ol>
	</li>
	<li>Once the skull is positioned in the stereotaxic device, determine and record the stereotactic coordinates of the bregma and lambda skull sutures along the anteroposterior (AP), medial lateral (ML), and dorsal ventral (DV) axes.</li>
	<li>Identify the target point (right hippocampus) with the aid of a stereotaxic atlas. Then, find on the skull the target point coordinates relative to bregma (AP: -2.7 mm, ML: -2.7 mm) (Figure 2E).</li>
	<li>Using a surgical marker, mark four corners of a 1.4 mm x 2 mm imaging window on the skull with dots, centered around the target point directly above the hippocampus (Figure 2F), for subsequent craniotomy.</li>
	<li>Use the surgical marker to draw two additional dots: one over the top-left parietal bone (Figure 2G) and another over the right frontal bone (Figure 2H), to indicate the future placement of two bone screws (Figure 3A, item b) that will anchor the head-cap (Figure 3B, item m) to the skull.</li>
	<li>Use the surgical marker to draw three more dots indicating where the epidural EEG recording electrodes will be inserted: one dot positioned 1 mm to either side of the midline, and one additional dot on the midline positioned 1 mm behind Lambda (Figure 2E, Figure 4A and Figure 5B).</li>
</ol>
2. Head-Cap installation
<ol>
	<li>Using a 0.7 mm diameter burr (Figure 3C), carefully drill two small holes in the skull, at the dot positions indicating the bone screws placement (see step 1.15 above). Then, take two bone screws (stainless steel-slotted, length: 4 mm; diameter: 0.85 mm) previously disinfected with ethanol 70-80%, and screw them to the skull for additional head-cap stability (Figure 2I-J and Figure 3A, item b).
	<ol>
		<li>Ensure that the screw tips do not protrude beyond the bottom of the bone into the skull cavity, risking damage to the brain. Note that one of the two bone screws will be anterior to the two head-cap screws, and the other posterior to them (Figure 2G-J).</li>
	</ol>
	</li>
	<li>Squirt saline over the skull to cool it down and clean it. Then dry the skull completely and apply a thin layer of cyanoacrylate around the screws.</li>
	<li>Use a custom alignment piece (Figure 3A, item c) clamped to the microdrive mounted to the stereotaxic device (Figure 2F-J and Figure 3A, item d), so that the position of the head-cap is aligned along the midline, with the anterior ridge overlying bregma. This custom stainless-steel piece is L-shaped (angled at 90&#176;), with two holes separated by 4 mm (Figure 2F-J and Figure 3A, item c).
	<ol>
		<li>Use the stereotaxic manipulator to position the L-shaped alignment piece, with two Fillister Head Slotted drive screws attached (M1.6 x 0.35 Metric Coarse, 12 mm Length; Figure 3A, item a) over the bregma, until the base of the screws lies flat on the skull, being careful to not exert pressure on the skull (Figure 2I-J).</li>
	</ol>
	</li>
	<li>Apply cyanoacrylate followed by dental cement to attach the base of the screws to the skull. Be careful to not obstruct the imaging window reference marks over the hippocampus (Figure 2K).</li>
</ol>
3. Imaging window craniotomy surgery
<ol>
	<li>At each of the positions of the craniotomy reference marks that indicate the corners of the imaging window (see step 1.14 above), drill a 0.7 mm diameter burr hole, followed by gently delineating the periphery of the 1.4 mm x 2 mm craniotomy window with the drill, with iteratively deeper cuts (Figure 2L).</li>
	<li>Take care to not to drill too deeply or with too much force on the skull, which is thin and fragile, having a thickness of approximately 300 &#181;m. Once the bone flap is complete, pry the loose flap up with the tip of a scalpel, being mindful to not damage the underlying dura mater.</li>
	<li>Cover the open window with bone wax (Figure 2M).</li>
	<li>Using a 0.9 mm diameter burr, drill three holes for future placement of the epidural EEG electrodes (see step 1.15 above) taking care to not to cut through the dura mater (Figure 2E,L).</li>
	<li>Cover the three holes with bone wax.</li>
	<li>Gently apply cyanoacrylate around the edges of the bone-wax covered imaging window and EEG holes, being careful to not seal the craniotomies (Figure 2L).</li>
	<li>Build a wall of dental cement that encompasses the imaging window and EEG holes and binds it to the previously cemented head-cap (Figure 2M and Figure 3B, item m).</li>
	<li>Let the cement dry for at least 10 minutes.</li>
	<li>Close any loose wound margins with simple interrupted sutures, using 4-0 or 5-0 black braided silk (Figure 2N and Figure 3C).</li>
</ol>
4. Post-surgery
<ol>
	<li>Use a cotton-tipped applicator to apply lidocaine ointment around the exposed skin to deaden sensation around the wound&#8217;s edge.</li>
	<li>Use a sterile cotton-tipped applicator to apply Neosporin to the exposed skin.</li>
	<li>Remove the animal from the stereotaxic frame (Figure 2N).</li>
	<li>Inject ketoprofen (5 mg/kg) IP or IM for post-surgical analgesia.</li>
	<li>Observe the animal until it is alert and moving (this will usually occur within minutes).</li>
	<li>Place the animal in a warm cage and continue to monitor its recovery until it drinks or eats and is ready to transfer to animal housing.</li>
	<li>Check on the animal at least once daily for approximately one week after the surgery, watching for any signs of listless behavior and/or inadequate eating/drinking (Figure 2O).</li>
</ol>
5. Dye injection and EEG recording
<ol>
	<li>Wait a minimum of one day after the head-cap implantation surgery to initiate the recordings.</li>
	<li>Place the mouse in a foam-restraining device molded to its body, within the stereotaxic frame (Figure 3B, items n &#38; j).</li>
	<li>Secure the head-cap (Figure 3B-I, item m) to a custom mounting bar which is attached to the stereotaxic frame (Figure 3A,B, items e &#38; j, and Figure 4A). The long section of item e (Figure 3A,B) should have a length between 9.4 &#8211; 13 mm. Fix item f (a mounting plate with dimensions 1.5 cm x 0.5 cm, with two holes separated 4 mm from center to center) to item e (Figure 3A,B, items e, f and k). This alignment system will position the awake mouse securely to the stereotaxic frame throughout the imaging sessions (Figure 4).</li>
	<li>Insert the epidural EEG recording electrodes (Figure 5A), placing the ground electrodes (white wires) within the posterior central hole, and the recording electrodes (red wires) in the anterior left and right holes (Figure 2E, Figure 4E and Figure 5E). Position each of the wires at an L-shape between the skull and the dura mater (Figure 5C) to optimize electrical contact.</li>
	<li>Inject the tail vein with green fluorescein lysine-fixable dextran (0.2 mL/25 g body wt. of Fluorescein 5% w/w), to visualize blood flow within the hippocampus.
	<ol>
		<li>To enhance dilation of the central vein of the mouse&#8217;s tail previous to injection, use one or more of the following strategies: a) Apply 70% ethanol to the central vein using a cotton-tipped applicator; b) Dip the tail into warm water (35-40 &#176;C) for 1-3 minutes; c) Expose the tail to a heating lamp for a few minutes.</li>
		<li>Secure the mouse&#8217;s tail with two fingers and locate the central vein, which lies immediately below the skin. Insert the needle (ultrafine insulin syringe with integrated needle 30G), with the bevel up, into the vein, approximately halfway to two thirds from the base of the tail. Keep the needle parallel to the tail during the injection.</li>
		<li>Inject the dye slowly and steadily, taking care to avoid any changes in the position of the needle or tail. 
		NOTE: There should be no resistance upon depressing the syringe plunger.</li>
		<li>After completing the injection, withdraw the needle and apply gentle pressure to the puncture site to seal the vessel.</li>
		<li>Allow clotting to occur.</li>
	</ol>
	</li>
</ol>
6. In vivo fiber-optic-bundle-coupled pre-clinical confocal laser-scanning endomicroscopy (pCLE)
<ol>
	<li>Directly following the tail vein injection (step 5.5.3 above), clamp the 300 &#181;m beveled fiber-optic bundle (containing 5000&#8211;7000 3 &#181;m fibers in each bundle) in the downward orientation to the mobile arm of the robotic stereotaxic drive.</li>
	<li>Remove the bone wax from the craniotomy.</li>
	<li>Remove the dura, using the bent tip of a syringe needle.</li>
	<li>Using the robotic positioning system, first move the fiber objective to the appropriate anterior-posterior and left-right stereotaxic location, with the tip zeroed in the z-axis on the surface of the cortex.</li>
	<li>Initiate the EEG recording (Figure 5D).</li>
	<li>Initiate confocal laser-scanning of the back plane of the fiber-objective, followed directly by descending the objective slowly in the z-axis using the robotic microdrive&#8217;s z-controls, until it reaches the target depth within the brain (Figure 4C). View the imaging results in the microscopic software while descending.
	<ol>
		<li>If the fiber tip is within the target X-Y-Z recording region, and two or more vessels enter the imaging window&#8217;s FOV, stop the descent and initiate video recordings. Obtain live full-field time-lapse movies of the blood flow at 11.7 Hz using Cellvizio Lab&#8217;s imaging software. (Higher speeds may be obtained with a smaller FOV). Recordings may take place for 4-5 hours at a time, in consecutive days if needed.</li>
	</ol>
	</li>
	<li>Using a 10 mL syringe with beveled silicon tubing fixed to the tip of the needle, feed the animal during the recordings by periodically providing paste made with mice pellets ground with water.</li>
	<li>At the end of the imaging session, terminate the recordings.</li>
	<li>Gently remove the EEG leads and clean them with Tergazyme.</li>
	<li>Slowly remove the fiber-optic bundle from the animal&#8217;s brain by reversing the fiber along the z-axis of entry.</li>
	<li>Release the animal from the head post and body restraints.</li>
	<li>If appropriate, follow the necessary euthanasia and tissue processing protocols to visualize blood vessels and to perform immunohistochemistry.</li>
	<li>If planning to record from the same animal in a future session, cover the imaging window with bone wax or silastic.</li>
</ol>

<ol>
	<li>Denk, 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=Anatomical+and+functional+imaging+of+neurons+using+2-photon+laser+scanning+microscopy.">Anatomical and functional imaging of neurons using 2-photon laser scanning microscopy.</a> Journal of Neuroscience Methods. 54 (2), 151-162 (1994).</li><li>Kleinfeld, D., Mitra, P. P., Helmchen, F., Denk, W. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Fluctuations+and+stimulus-induced+changes+in+blood+flow+observed+in+individual+capillaries+in+layers+2+through+4+of+rat+neocortex.">Fluctuations and stimulus-induced changes in blood flow observed in individual capillaries in layers 2 through 4 of rat neocortex.</a> Proceedings of the National Academy of Sciences of the United States of America. 95, 15741-15746 (1998).</li><li>Helmchen, F., Fee, M. S., Tank, D. W., Denk, W. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+miniature+head-mounted+two-photon+microscope.+High-resolution+brain+imaging+in+freely+moving+animals.">A miniature head-mounted two-photon microscope. High-resolution brain imaging in freely moving animals.</a> Neuron. 31, 903 (2001).</li><li>Chaigneau, E., Oheim, M., Audinat, E., Charpak, S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Two-photon+imaging+of+capillary+blood+flow+in+olfactory+bulb+glomeruli.">Two-photon imaging of capillary blood flow in olfactory bulb glomeruli.</a> Proceedings of the National Academy of Sciences of the United States of America. 100, 13081-13086 (2003).</li><li>Larson, D. R., 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=Water-soluble+quantum+dots+for+multiphoton+fluorescence+imaging+in+vivo.">Water-soluble quantum dots for multiphoton fluorescence imaging in vivo.</a> Science. 300, 1434-1436 (2003).</li><li>Hirase, H., Creso, J., Singleton, M., Bartho, P., Buzsaki, G. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Two-photon+imaging+of+brain+pericytes+in+vivo+using+dextran-conjugated+dyes.">Two-photon imaging of brain pericytes in vivo using dextran-conjugated dyes.</a> Glia. 46, 95-100 (2004).</li><li>Hirase, H., Creso, J., Buzsaki, G. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Capillary+level+imaging+of+local+cerebral+blood+flow+in+bicuculline-induced+epileptic+foci.">Capillary level imaging of local cerebral blood flow in bicuculline-induced epileptic foci.</a> Neuroscience. 128, 209-216 (2004).</li><li>Schaffer, C. B., 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=Two-photon+imaging+of+cortical+surface+microvessels+reveals+a+robust+redistribution+in+blood+flow+after+vascular+occlusion.">Two-photon imaging of cortical surface microvessels reveals a robust redistribution in blood flow after vascular occlusion.</a> PLoS Biology. 4, 22 (2006).</li><li>Leal-Campanario, R., 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=Abnormal+Capillary+Vasodynamics+Contribute+to+Ictal+Neurodegeneration+in+Epilepsy.">Abnormal Capillary Vasodynamics Contribute to Ictal Neurodegeneration in Epilepsy.</a> Scientific Reports. 7, 43276 (2017).</li><li>Peppiatt, C. M., Howarth, C., Mobbs, P., Attwell, D. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Bidirectional+control+of+CNS+capillary+diameter+by+pericytes.">Bidirectional control of CNS capillary diameter by pericytes.</a> Nature. 443, 700-704 (2006).</li><li>Yemisci, M., 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=Pericyte+contraction+induced+by+oxidative-nitrative+stress+impairs+capillary+reflow+despite+successful+opening+of+an+occluded+cerebral+artery.">Pericyte contraction induced by oxidative-nitrative stress impairs capillary reflow despite successful opening of an occluded cerebral artery.</a> Nature Medicine. 15, 1031-1037 (2009).</li><li>Fernandez-Klett, F., Offenhauser, N., Dirnagl, U., Priller, J., Lindauer, U. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Pericytes+in+capillaries+are+contractile+in+vivo,+but+arterioles+mediate+functional+hyperemia+in+the+mouse+brain.">Pericytes in capillaries are contractile in vivo, but arterioles mediate functional hyperemia in the mouse brain.</a> Proceedings of the National Academy of Sciences of the United States of America. 107, 22290-22295 (2010).</li><li>Leal-Campanario, R., Alarcon-Martinez, L., Martinez-Conde, S., Calhoun, M., Macknik, S., Mouton, P. R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Blood+Flow+Analysis+in+Epilepsy+Using+a+Novel+Stereological+Approach.">Blood Flow Analysis in Epilepsy Using a Novel Stereological Approach.</a> Neurostereology: Unbiased Stereology of Neural Systems. , (2013).</li><li>Smart, S. L., 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=Deletion+of+the+K(V)1.1+potassium+channel+causes+epilepsy+in+mice.">Deletion of the K(V)1.1 potassium channel causes epilepsy in mice.</a> Neuron. 20, 809-819 (1998).</li><li>Zuberi, S. M., 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+novel+mutation+in+the+human+voltage-gated+potassium+channel+gene+(Kv1.1)+associates+with+episodic+ataxia+type+1+and+sometimes+with+partial+epilepsy.">A novel mutation in the human voltage-gated potassium channel gene (Kv1.1) associates with episodic ataxia type 1 and sometimes with partial epilepsy.</a> Brain. 122, 817-825 (1999).</li></ol>

We have nothing to disclose.

We developed a head-cap restraint system for simultaneous electrophysiological and fiber-optic pCLE experiments in awake mice, reducing potential response contamination due to anesthetic drugs. The head-cap and mounting apparatus are straightforward to construct and are reusable for chronic awake-behaving imaging experiments. We checked the quality of the recordings against the gold-standard for in vivo microscopic blood flow imaging, TPLSM.
Proficient surgical skills are necessary to implemen...

In contrast with other microscopic imaging methods1,2,3,4,5,6,7,8, in vivo fiber-optic-based confocal microscopy allows the measurement of blood flow dynamics in any brain region, at any depth, at high speed (up to 240 Hz depending on field-of-view size9). A fiber-optic probe enables in vivo confocal laser scanning imaging at 3 &#181;m resolution because the tip of the probe (a lens-less objective made up of a bundle of 5000-6000 3 &#181;m diameter individual fibers) can be positioned with a microelectrode&#8217;s accuracy, within 15 &#181;m of the fluorescent target of interest. As with in vivo two-photon imaging, fluorophores must be previously introduced into the imaging target. For example, fluorescein dextran (or quantum dots) may be injected into the vasculature, or genetically-encoded fluorescent proteins can be transfected into cells, or fluorescent dyes such as Oregon Green BAPTA-1 can be bulk-loaded into cells, prior to imaging.
Recent research using these techniques has found that mural cell motor activity leading to ictal capillary vasospasms&#8212;sudden constrictions that occur at the position of the mural cells during seizures9&#8212;can contribute to neurodegeneration in the ictal hippocampus9. Whereas previous imaging studies showed in vitro and in vivo pericyte constrictions connected to drug applications6,7,10,11,12, Leal-Campanario et al. found the first evidence of in vivo spontaneous capillary constrictions in the murine brain. To establish relevance to human temporal lobe epilepsy, they studied male (P30-40 old) knockout (KO) Kv1.1 (kcna1-null) mice14,15 (JAX stock #003532), a genetic model of human episodic ataxia type 115. Pericytes drove both pathological and physiological hippocampal mural vasoconstrictions9 in the spontaneously epileptic animals and their wild-type (WT) littermates. These observations were replicated in WT animals rendered epileptic with kainic-acid, thereby indicating their generalization to other forms of epilepsy. Leal-Campanario et al moreover determined, using novel stereological microscopy approaches, that apoptotic&#8212;but not healthy&#8212;neurons in epileptic animals were spatially coupled to the hippocampal microvasculature. Because excitotoxicity has no known spatial association to the vasculature, this result indicated that abnormal capillary vasospasmic ischemia-induced hypoxia contributes to neurodegeneration in epilepsy. Figure 1 shows a schematic of the general setup.

<table>
	<tbody>
		<tr>
			<td>0.7 mm diameter burr</td>
			<td>Fine Science Tools</td>
			<td>19007-07</td>
			<td>For Screws No. 19010-00</td>
		</tr>
		<tr>
			<td>0.9 mm diameter burr</td>
			<td>Fine Science Tools</td>
			<td>19007-09</td>
			<td></td>
		</tr>
		<tr>
			<td>ASEPTICO AEU-12C TORQUE PLUS</td>
			<td>from Handpiece solution</td>
			<td>AEU12C</td>
			<td></td>
		</tr>
		<tr>
			<td>Bull dog serrifine clump</td>
			<td>Fine Science Tools</td>
			<td>18050-28</td>
			<td></td>
		</tr>
		<tr>
			<td>CellVizio dual band</td>
			<td>Mauna Kea Technologies</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>CellVizio single band</td>
			<td>Mauna Kea Technologies</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Confocal Microprobe 300 microns (Serie S)</td>
			<td>Mauna Kea Technologies</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Custom-made alignment piece</td>
			<td>L-shaped (angled at 90 deg) and made of stainless steel with two holes drilled on it, with a 4 mm separation from center to center</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Custom-made mounting bar</td>
			<td>The long section piece of the mounting bar should be between 9.4 - 13mm. Fixed to this piece of the mounting bar, position a stainless-steel plate 1.5 cm long and 0.5 cm wide that has two holes drilled separated 4 mm from center to center, the same distance that the L-shaped alignment piece.</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Cyanoacrylate adhesive-Super Glue</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Dumont forceps #5</td>
			<td>Fine Science Tools</td>
			<td>11252-20</td>
			<td></td>
		</tr>
		<tr>
			<td>DuraLay Inlay Resin &#8211; Standard Package</td>
			<td>Reliance Dental Mfg Co.</td>
			<td>602-7395 (from patterson dental)</td>
			<td></td>
		</tr>
		<tr>
			<td>Fillister Head, Slotted Drive, M1.6x0.35 Metric Coarse, 12mm Length Under Head, Machine Screw</td>
			<td>MSC industrial direct co.</td>
			<td>2834117</td>
			<td></td>
		</tr>
		<tr>
			<td>Fine Point scissor</td>
			<td>Fine Science Tools</td>
			<td>14090-09</td>
			<td></td>
		</tr>
		<tr>
			<td>Fluorescein 5% w/w lysine-fixable dextran (2MD)</td>
			<td>Invitrogen, USA</td>
			<td>D7137</td>
			<td></td>
		</tr>
		<tr>
			<td>Halsey smooth needle holder</td>
			<td>Fine Science Tools</td>
			<td>12001-13</td>
			<td></td>
		</tr>
		<tr>
			<td>Kalt suture needle 3/8 curved</td>
			<td>Fine Science Tools</td>
			<td>12050-03</td>
			<td></td>
		</tr>
		<tr>
			<td>lab standard stereotaxic, rat and mouse</td>
			<td>Stoelting Co. 51704</td>
			<td>51670</td>
			<td></td>
		</tr>
		<tr>
			<td>Methocel 2%</td>
			<td>Omnivision GmbH</td>
			<td>PZN: 04682367</td>
			<td>Eye ointment to prevent dryness.</td>
		</tr>
		<tr>
			<td>Mouse Temperature controller, probe (YSI-451), small heating pad-TC-1000 Mouse</td>
			<td>CWE Inc.</td>
			<td>08-13000</td>
			<td></td>
		</tr>
		<tr>
			<td>PhysioTel F20-EET transmitters</td>
			<td>DSI</td>
			<td>270-0124-001</td>
			<td></td>
		</tr>
		<tr>
			<td>Robot Stereotaxic, Manipulator Arm, ADD-ON, 3 Axis, LEFT</td>
			<td>Stoelting Co.C13</td>
			<td>51704</td>
			<td></td>
		</tr>
		<tr>
			<td>Sel-Tapping bone screws</td>
			<td>Fine Science Tools</td>
			<td>19010-10</td>
			<td></td>
		</tr>
		<tr>
			<td>Standard Ear Bars and Rubber Tips for Mouse Stereotaxic</td>
			<td>Stoelting Co</td>
			<td>51648</td>
			<td></td>
		</tr>
		<tr>
			<td>Suture Thread - Braided Silk/Size 4/0</td>
			<td>Fine Science Tools</td>
			<td>18020-40</td>
			<td></td>
		</tr>
		<tr>
			<td>Tissue separating microspatula</td>
			<td>Fine Science Tools</td>
			<td>10091-121</td>
			<td></td>
		</tr>
	</tbody>
</table>

in vivo fiber-coupled pre-clinical confocal laser-scanning endomicroscopy (pcle) of hippocampal capillaries in awake mice

The goal of this protocol is to describe fiber-optic-bundle-coupled pre-clinical confocal laser-scanning endomicroscopy (pCLE) in its specific application to elucidate capillary blood flow effects during seizures, driven by mural cells. In vitro and in vivo cortical imaging have shown that capillary constrictions driven by pericytes can result from functional local neural activity, as well as from drug application, in healthy animals. Here, a protocol is presented on how to use pCLE to determine the role of microvascular dynamics in neural degeneration in epilepsy, at any tissue depth (specifically in the hippocampus). We describe a head restraint technique that has been adapted to record pCLE in awake animals, to address potential side-effects of anesthetics on neural activity. Using these methods, electrophysiological and imaging recordings can be conducted over several hours in deep neural structures of the brain.

We developed these methods to assess whether abnormal pericyte-driven capillary vasospasms in the hippocampus&#8212;occurring as the result of seizures&#8212;could cause frank hypoxia that contributes to cell death in the ictal focus9,13.
The development of the head-cap and its proper installation afforded high-stability in the recordings, allowing simultaneous recording of EEG and blood flow deep in the hippocampus of wild-type and ep...

Watch this Scientific Journal Video about In Vivo Fiber-Coupled Pre-Clinical Confocal Laser-scanning Endomicroscopy pCLE of Hippocampal Capillaries in Awake Mice at JoVE.com

In Vivo Fiber-Coupled Pre-Clinical Confocal Laser-scanning Endomicroscopy pCLE of Hippocampal Capillaries in Awake Mice

Whereas multiphoton imaging is only effective at limited depths from the tissue&#8217;s surface, it is possible to achieve 3 &#181;m resolution imaging at any depth via pCLE. Here, we present a protocol to conduct pCLE imaging to measure microvascular dynamics in the hippocampus of ictal and wild-type mice.

In Vivo Fiber-Coupled Pre-Clinical Confocal Laser-scanning Endomicroscopy (pCLE) of Hippocampal Capillaries in Awake Mice

The goal of this protocol is to describe fiber-optic-bundle-coupled pre-clinical confocal laser-scanning endomicroscopy (pCLE) in its specific application ...

in-vivo-fiber-coupled-pre-clinical-confocal-laser-scanning

Research

JoVE Journal

Neuroscience

837 Views.  Universidad Pablo de Olavide. Whereas multiphoton imaging is only effective at limited depths from the tissue’s surface, it is possible to achieve 3 µm resolution imaging at any depth via pCLE. Here, we present a protocol to conduct pCLE imaging to measure microvascular dynamics in the hippocampus of ictal and wild-type mice.

Video: In Vivo Fiber-Coupled Pre-Clinical Confocal Laser-scanning Endomicroscopy pCLE of Hippocampal Capillaries in Awake Mice

Transfection of Mouse Retinal Ganglion Cells by in vivo Electroporation

The targeting and refinement of RGC projections to the midbrain is a popular and powerful model system for studying how precise patterns of neural connectivity form during development. In mice, retinofugal projections are arranged in a topographic manner and form eye-specific layers in the Lateral Geniculate Nucleus (dLGN) of the thalamus and the Superior Colliculus (SC). The development of these precise patterns of retinofugal projections has typically been studied by labeling populations of RGCs with fluorescent dyes and tracers, such as horseradish peroxidase1-4. However, these methods are too coarse to provide insight into developmental changes in individual RGC axonal arbor morphology that are the basis of retinotopic map formation. They also do not allow for the genetic manipulation of RGCs.
Recently, electroporation has become an effective method for providing precise spatial and temporal control for delivery of charged molecules into the retina5-11. Current retinal electroporation protocols do not allow for genetic manipulation and tracing of retinofugal projections of a single or small cluster of RGCs in postnatal mice. It has been argued that postnatal in vivo electroporation is not a viable method for transfecting RGCs since the labeling efficiency is extremely low and hence requires targeting at embryonic ages when RGC progenitors are undergoing differentiation and proliferation6. 
In this video we describe an in vivo electroporation protocol for targeted delivery of genes, shRNA, and fluorescent dextrans to murine RGCs postnatally. This technique provides a cost effective, fast and relatively easy platform for efficient screening of candidate genes involved in several aspects of neural development including axon retraction, branching, lamination, regeneration and synapse formation at various stages of circuit development. In summary we describe here a valuable tool which will provide further insights into the molecular mechanisms underlying sensory map development.

Transfection of Mouse Retinal Ganglion Cells by in vivo Electroporation

We demonstrate an in vivo electroporation protocol for transfecting single or small clusters of retinal ganglion cells (RGCs) and other retinal cell types in postnatal mice over a wide range of ages. The ability to label and genetically manipulate postnatal RGCs in vivo is a powerful tool for developmental studies.

The targeting and refinement of RGC projections to the midbrain is a popular and powerful model system for studying how precise patterns of neural ...

Mapping Inhibitory Neuronal Circuits by Laser Scanning Photostimulation

Inhibitory neurons are crucial to cortical function. They comprise about 20% of the entire cortical neuronal population and can be further subdivided into diverse subtypes based on their immunochemical, morphological, and physiological properties1-4. Although previous research has revealed much about intrinsic properties of individual types of inhibitory neurons, knowledge about their local circuit connections is still relatively limited3,5,6. Given that each individual neuron's function is shaped by its excitatory and inhibitory synaptic input within cortical circuits, we have been using laser scanning photostimulation (LSPS) to map local circuit connections to specific inhibitory cell types. Compared to conventional electrical stimulation or glutamate puff stimulation, LSPS has unique advantages allowing for extensive mapping and quantitative analysis of local functional inputs to individually recorded neurons3,7-9. Laser photostimulation via glutamate uncaging selectively activates neurons perisomatically, without activating axons of passage or distal dendrites, which ensures a sub-laminar mapping resolution. The sensitivity and efficiency of LSPS for mapping inputs from many stimulation sites over a large region are well suited for cortical circuit analysis.
Here we introduce the technique of LSPS combined with whole-cell patch clamping for local inhibitory circuit mapping. Targeted recordings of specific inhibitory cell types are facilitated by use of transgenic mice expressing green fluorescent proteins (GFP) in limited inhibitory neuron populations in the cortex3,10, which enables consistent sampling of the targeted cell types and unambiguous identification of the cell types recorded. As for LSPS mapping, we outline the system instrumentation, describe the experimental procedure and data acquisition, and present examples of circuit mapping in mouse primary somatosensory cortex. As illustrated in our experiments, caged glutamate is activated in a spatially restricted region of the brain slice by UV laser photolysis; simultaneous voltage-clamp recordings allow detection of photostimulation-evoked synaptic responses. Maps of either excitatory or inhibitory synaptic input to the targeted neuron are generated by scanning the laser beam to stimulate hundreds of potential presynaptic sites. Thus, LSPS enables the construction of detailed maps of synaptic inputs impinging onto specific types of inhibitory neurons through repeated experiments. Taken together, the photostimulation-based technique offers neuroscientists a powerful tool for determining the functional organization of local cortical circuits.

This paper introduces an approach of combining laser scanning photostimulation with whole cell recordings in transgenic mice expressing GFP in limited inhibitory neuron populations. The technique allows for extensive mapping and quantitative analysis of local synaptic circuits of specific inhibitory cortical neurons.

Inhibitory neurons are crucial to cortical function. They comprise about 20% of the entire cortical neuronal population and can be further subdivided into ...

Functional Neuroimaging Using Ultrasonic Blood-brain Barrier Disruption and Manganese-enhanced MRI

Although mice are the dominant model system for studying the genetic and molecular underpinnings of neuroscience, functional neuroimaging in mice remains technically challenging. One approach, Activation-Induced Manganese-enhanced MRI (AIM MRI), has been used successfully to map neuronal activity in rodents 1-5. In AIM MRI, Mn2+ acts a calcium analog and accumulates in depolarized neurons 6,7. Because Mn2+ shortens the T1 tissue property, regions of elevated neuronal activity will enhance in MRI. Furthermore, Mn2+ clears slowly from the activated regions; therefore, stimulation can be performed outside the magnet prior to imaging, enabling greater experimental flexibility. However, because Mn2+ does not readily cross the blood-brain barrier (BBB), the need to open the BBB has limited the use of AIM MRI, especially in mice.
One tool for opening the BBB is ultrasound. Though potentially damaging, if ultrasound is administered in combination with gas-filled microbubbles (i.e., ultrasound contrast agents), the acoustic pressure required for BBB opening is considerably lower. This combination of ultrasound and microbubbles can be used to reliably open the BBB without causing tissue damage 8-11.
Here, a method is presented for performing AIM MRI by using microbubbles and ultrasound to open the BBB. After an intravenous injection of perflutren microbubbles, an unfocused pulsed ultrasound beam is applied to the shaved mouse head for 3 minutes. For simplicity, we refer to this technique of BBB Opening with Microbubbles and UltraSound as BOMUS 12. Using BOMUS to open the BBB throughout both cerebral hemispheres, manganese is administered to the whole mouse brain. After experimental stimulation of the lightly sedated mice, AIM MRI is used to map the neuronal response.
To demonstrate this approach, herein BOMUS and AIM MRI are used to map unilateral mechanical stimulation of the vibrissae in lightly sedated mice 13. Because BOMUS can open the BBB throughout both hemispheres, the unstimulated side of the brain is used to control for nonspecific background stimulation. The resultant 3D activation map agrees well with published representations of the vibrissae regions of the barrel field cortex 14. The ultrasonic opening of the BBB is fast, noninvasive, and reversible; and thus this approach is suitable for high-throughput and/or longitudinal studies in awake mice.

A technique is described for broadly opening the blood-brain barrier in the mouse using microbubbles and ultrasound. Using this technique, manganese can be administered to the mouse brain. Because manganese is an MRI contrast agent that accumulates in depolarized neurons, this approach enables imaging of neuronal activity.

Although mice are the dominant model system for studying the genetic and molecular underpinnings of neuroscience, functional neuroimaging in mice remains ...

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

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

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

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

Laser-scanning Photostimulation of Optogenetically Targeted Forebrain Circuits

The sensory forebrain is composed of intricately connected cell types, of which functional properties have yet to be fully elucidated. Understanding the interactions of these forebrain circuits has been aided recently by the development of optogenetic methods for light-mediated modulation of neuronal activity. Here, we describe a protocol for examining the functional organization of forebrain circuits&#160;in vitro using laser-scanning photostimulation of channelrhodopsin, expressed optogenetically via viral-mediated transfection. This approach also exploits the utility of cre-lox recombination in transgenic mice to target expression in specific neuronal cell types. Following transfection, neurons are physiologically recorded in slice preparations using whole-cell patch clamp to measure their evoked responses to laser-scanning photostimulation of channelrhodopsin expressing fibers. This approach enables an assessment of functional topography and synaptic properties. Morphological correlates can be obtained by imaging the neuroanatomical expression of channelrhodopsin expressing fibers using confocal microscopy of the live slice or post-fixed tissue. These methods enable functional investigations of forebrain circuits that expand upon more conventional approaches.

We describe a method for characterizing the functional topography and synaptic properties of forebrain circuits using an optogenetic approach to photostimulate neuronal populations&#160;in vitro.

The sensory forebrain is composed of intricately connected cell types, of which functional properties have yet to be fully elucidated. Understanding the ...

A Simple and Inexpensive Method for Determining Cold Sensitivity and Adaptation in Mice

Cold hypersensitivity is a serious clinical problem, affecting a broad subset of patients and causing significant decreases in quality of life. The cold plantar assay allows the objective and inexpensive assessment of cold sensitivity in mice, and can quantify both analgesia and hypersensitivity. Mice are acclimated on a glass plate, and a compressed dry ice pellet is held against the glass surface underneath the hindpaw. The latency to withdrawal from the cooling glass is used as a measure of cold sensitivity.
Cold sensation is also important for survival in regions with seasonal temperature shifts, and in order to maintain sensitivity animals must be able to adjust their thermal response thresholds to match the ambient temperature. The Cold Plantar Assay (CPA) also allows the study of adaptation to changes in ambient temperature by testing the cold sensitivity of mice at temperatures ranging from 30 &#176;C to 5 &#176;C. Mice are acclimated as described above, but the glass plate is cooled to the desired starting temperature using aluminum boxes (or aluminum foil packets) filled with hot water, wet ice, or dry ice. The temperature of the plate is measured at the center using a filament T-type thermocouple probe. Once the plate has reached the desired starting temperature, the animals are tested as described above.
This assay allows testing of mice at temperatures ranging from innocuous to noxious. The CPA yields unambiguous and consistent behavioral responses in uninjured mice and can be used to quantify both hypersensitivity and analgesia. This protocol describes how to use the CPA to measure cold hypersensitivity, analgesia, and adaptation in mice.

The Cold Plantar Assay (CPA) measures cold responsiveness between 30 &#176;C and 5 &#176;C, and can also measure cold adaptation. This protocol describes how to use the CPA to measure cold hypersensitivity, analgesia, and adaptation in mice.

Cold hypersensitivity is a serious clinical problem, affecting a broad subset of patients and causing significant decreases in quality of life.  The cold ...

Simultaneous ex vivo Functional Testing of Two Retinas by in vivo Electroretinogram System

An In vivo electroretinogram (ERG) signal is composed of several overlapping components originating from different retinal cell types, as well as noise from extra-retinal sources. Ex vivo ERG provides an efficient method to dissect the function of retinal cells directly from an intact isolated retina of animals or donor eyes. In addition, ex vivo ERG can be used to test the efficacy and safety of potential therapeutic agents on retina tissue from animals or humans. We show here how commercially available in vivo ERG systems can be used to conduct ex vivo ERG recordings from isolated mouse retinas. We combine the light stimulation, electronic and heating units of a standard in vivo system with custom-designed specimen holder, gravity-controlled perfusion system and electromagnetic noise shielding to record low-noise ex vivo ERG signals simultaneously from two retinas with the acquisition software included in commercial in vivo systems. Further, we demonstrate how to use this method in combination with pharmacological treatments that remove specific ERG components in order to dissect the function of certain retinal cell types.

Simultaneous ex vivo Functional Testing of Two Retinas by in vivo Electroretinogram System

Ex vivo ERG can be used to record electrical activity of retinal cells directly from isolated intact retinas of animals or humans. We demonstrate here how common in vivo ERG systems can be adapted for ex vivo ERG recordings in order to dissect the electrical activity of retinal cells.

An In vivo electroretinogram (ERG) signal is composed of several overlapping components originating from different retinal cell types, as well as noise ...

Protocol for Three-dimensional Confocal Morphometric Analysis of Astrocytes

As glial cells in the brain, astrocytes have diverse functional roles in the central nervous system. In the presence of harmful stimuli, astrocytes modify their functional and structural properties, a condition called reactive astrogliosis. Here, a protocol for assessment of the morphological properties of astrocytes is presented. This protocol includes quantification of 12 different parameters i.e. the surface area and volume of the tissue covered by an astrocyte (astrocyte territory), the entire astrocyte including branches, cell body, and nucleus, as well as total length and number of branches, the intensity of fluorescence immunoreactivity of antibodies used for astrocyte detection, and astrocyte density (number/1,000 &#181;m2). For this purpose three-dimensional (3D) confocal microscopic images were created, and 3D image analysis software such as Volocity 6.3 was used for measurements. Rat brain tissue exposed to amyloid beta1-40 (A&#946;1-40) with or without a therapeutic&#160;intervention was used to present the method. This protocol can also be used for 3D morphometric analysis of other cells from either in vivo or in vitro conditions.

Astrocytes in the CNS change their functional and structural properties in response to harmful stimuli. This report presents a protocol for assessment of three-dimensional astrocyte morphology in diseased conditions or after therapeutic interventions.

As glial cells in the brain, astrocytes have diverse functional roles in the central nervous system. In the presence of harmful stimuli, astrocytes modify ...

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 ...

Optogenetics Identification of a Neuronal Type with a Glass Optrode in Awake Mice

It is a major concern in neuroscience how different types of neurons work in neural circuits. Recent advances in optogenetics have enabled the identification of the neuronal type in in vivo electrophysiological experiments in broad brain regions. In optogenetics experiments, it is critical to deliver the light to the recording site. However, it is often hard to deliver the stimulation light to the deep brain regions from the brain&#39;s surface. Especially, it is difficult for the stimulation light to reach the deep brain regions when the optical transparency of the brain surface is low, as is often the case with recordings from awake animals. Here, we describe a method to record spike responses to the light from an awake mouse using a custom-made glass optrode. In this method, the light is delivered through the recording glass electrode so that it is possible to reliably stimulate the recorded neuron with light in the deep brain regions. This custom-made optrode system consists of accessible and inexpensive materials and is easy to assemble.

This work introduces a method to perform an optogenetic single-unit recording reliably from an awake mouse using a custom-made glass optrode.