The majority of the work and the personnel involved were supported by NIH grant NS060677 and partly by NIH grants MH099559 and MH104069 (to BSK). Some of the work was also supported by the CHDI Foundation.

All animal protocols were in accordance with the US National Institutes of Health Guide for the Care and Use of Laboratory Animals and were approved by the Institutional Animal Care and Use Committee at UCLA.
1.1) Prepare Micropipette and AAV2/5 Virus Loading
<ol>
	<li>Use fine borosilicate glass micropipettes for the injection of the virus. Pull the micropipette using a two-step pulling program with a vertical puller. Bevel the pipette at an angle of 40&#176; by using a pipette grinder. The pipette that is ideal for use will have a tip diameter of 20 - 40 &#956;m and a shank length of 6 - 7 mm. Autoclave the pipette and surgical instruments. The drill is sterilized by wiping with 70% ethanol.</li>
	<li>Store the virus AAV2/5-GfaABC1D.cyto-GCaMP3.SV40 (1.5 x 1013 gc/ml), at -80 &#176;C in 10 &#956;l aliquots. Just before surgery, take out the aliquots out of the freezer and store on ice until used for filling the micropipette.</li>
	<li>Fill the micropipette with virus using a syringe pump. Tightly connect one end of a piece of tubing, through an appropriately sized tubing compression fitting, to a glass syringe with a reinforced plunger. Attach the autoclaved micropipette to the other end of the piece of tubing through an appropriately sized removable needle compression fitting.</li>
	<li>Before loading the viral vector, remove air bubbles from the pumping system including tubing, syringe and micropipette by filling the system with mineral oil colored with Sudan red IV (~1 mg/50 ml) in a 1 ml syringe with a needle (27 G).</li>
	<li>Place a clean piece of parafilm under the glass micropipette, and dispense 5 - 10 &#956;l of the viral vector onto the parafilm.</li>
	<li>Suck the viral vector by moving the plunger of the syringe backward at 0.5 &#956;l/min, and mark the boundary between the vector and the oil on the glass pipette.</li>
</ol>
1.2) Anesthesia, Hair Cutting, Head Fixation and Preparation for Craniotomy
<ol>
	<li>Put a mouse (P49-63, C57BL/6) into the chamber filled with N2O and O2, and 5% isoflurane. Assess the depth of anesthesia by loss of purposeful movement and a slowed respiratory rate (~60 - 90/min). Cut hair on the head when the mouse is anesthetized.</li>
	<li>Fit the mouse in the stereotaxic frame, with its head secured by blunt ear bars and its nose placed into an anesthesia and ventilation system. Maintain continuous isoflurane at 2 - 3%. Apply artificial tears ointment to both eyes since mice lose their blink reflex under anesthesia.</li>
	<li>Administer 0.05 ml of Buprenorphine (0.1 mg/ml) by subcutaneous injection for pain relief.</li>
	<li>Clean surgical area with 10% povidone iodine and 70% alcohol three times, alternating between the two solutions and starting with povidone iodine. Start from the front and wipe toward the back to remove any already cut hairs.</li>
	<li>Make a skin incision on top of the head which starts between the eyes and ends between the ears.</li>
</ol>
1.3) Craniotomy and Microinjections
<ol>
	<li>Remove the periosteum by swiping with a cotton swab and then dry the surface of the skull with dental cotton balls. Make a mark around the injection site, and drill around the edge of the mark using a small steel burr powered by a high speed drill.</li>
	<li>Remove the bone and clean the surface with saline.</li>
	<li>Place the pipette into the left dorsal lateral striatum with the following coordinates (mm): anterior-posterior +0.8, medial-lateral: +2.0, dorsal-ventral from the pial surface: -2.4 13. Note that these coordinates mean that the tip of the microinjection needle sits just above the dorsolateral striatum (Figure 1A, B), meaning that it only slightly penetrates within this nucleus. Avoid damaging blood vessels. If the pipette happens to be right on a blood vessel, slightly adjust the coordinates by ~0.1 - 0.2 mm.</li>
	<li>Start injection with injection rate set at 0.2 &#956;l/min, and typically inject 1 to 1.5&#160;&#956;l of virus.</li>
	<li>Leave the pipette in place after the injection for 10 min, and then withdraw the pipette slowly.</li>
	<li>Finish by closing the surgical wound with continuous suture external nylon suture.</li>
</ol>
1.4) Recovery
<ol>
	<li>Put the mouse in a clean cage on a heating pad for 24 hr. Do not return a mouse that has undergone surgery to the company of other animals until fully recovered. Do not leave the mouse unattended until it has regained sufficient consciousness to maintain sternal recumbency.</li>
	<li>Add Trimethoprim Sulfamethoxiazole (5 ml per 500 ml water, containing 40 mg trimethoprim and 200 mg sulfamethoxiazole) in the water for a week. Administer Buprenorphine two times per day for up to 3 days after surgery.</li>
	<li>Maintain the mouse on a 12 hr light-dark cycle, fed and watered daily, and is weighted once per day up to 3 days after surgery. In these 3 days, any mouse with a loss of more than 10% body weight is considered as compromised, and will be euthanized instead of being subjected to experimentation.</li>
	<li>Separately, change the mouse cage twice per week, and monitor for general health at least once per day. Typically, the mouse is used within 2 - 3 weeks after the stereotaxic microinjections.</li>
</ol>
2. Acute Brain Slice Preparation for Confocal Ca2+ Imaging
2.1) Preparation of Cutting and Recording Solutions.
<ol>
	<li>Prepare cutting solution comprising (mM): 194 sucrose, 30 NaCl, 4.5 KCl, 10 D-glucose, 1 MgCl2, 1.2 NaH2PO4, and 26 NaHCO3, saturated with 95% O2 and 5% CO2.</li>
	<li>Prepare recording solution comprising (mM): 124 NaCl, 4.5 KCl, 1 MgCl2, 10 D-glucose, 2 CaCl2, 1.2 NaH2PO4, and 26 NaHCO3; pH 7.3 - 7.4, 290 - 295 mOsm, saturated with 95% O2 and 5% CO2. Fill the brain slice holding beaker with recording solution, and keep it at 34 &#176;C.</li>
</ol>
2.2) Slicing
<ol>
	<li>Two to three weeks after stereotaxic microinjections, deeply and terminally anesthetize the mouse, and then decapitate it.</li>
	<li>Extract the brain, and use a blade to remove the uninjected right hemisphere, and mount the left one onto the vibratome tray using super glue. Fill the vibratome tray with ice cold cutting buffer, and keep saturating it with 95% O2 and 5% CO2.</li>
	<li>Cut coronal or sagittal striatal brain slices at 300&#160;&#956;m thickness. Usually, 4 - 5 coronal, or 6 - 7 sagittal striatal slices can be collected.</li>
	<li>Transfer the slices to the slice holding beaker warmed at 34 &#176;C, and keep them there for 30 min before storing them at room temperature for subsequent recording.</li>
	<li>Incubate the brain slices in oxygenated recording buffer at room temperature for at least 30 min before [Ca2+]i imaging.</li>
</ol>
3. Immunohistochemistry (IHC)
<ol>
	<li>Two weeks after virus injection, perfuse the mouse transcardially with PBS followed by 10% formalin in PBS.</li>
	<li>Remove, post-fix overnight, and cryoprotect the brain in buffered 30% sucrose for at least 2 days.</li>
	<li>Prepare 40 &#956;m sections using a cryostat microtome.</li>
	<li>Rinse sections 3 times with PBS at 10 min interval.</li>
	<li>Incubate sections for 1 hr in PBS with 10% normal goat serum and with 0.5% Triton X-100.</li>
	<li>Incubate sections overnight at 4 &#176;C in primary antibody prepared in PBS with 0.5% Triton X-100. The primary antibodies used were as follows: chicken anti-GFP 1:1,000, mouse anti-S100&#946; 1:400, mouse anti-NeuN 1:600, mouse anti-glutamine synthetase 1:300.</li>
	<li>Rinse sections 3 times with PBS at 10 min intervals.</li>
	<li>Incubate sections with the secondary antibodies prepared in PBS with 10% normal goat serum for 2 hr at room temperature. The second antibodies were as follows: goat anti-mouse Alexa546 1:500, goat anti-chicken Alexa488 1:500.</li>
	<li>Mount the sections on glass slides, dry and apply several drops of antifade mounting medium, then slowly cover the sections with a glass coverslip. Store the slides at 4 &#176;C.</li>
	<li>Take images on a confocal microscope with a 40X oil immersion lens with a numerical aperture of 1.3.</li>
</ol>
4. Confocal [Ca2+]i Imaging
NOTE: [Ca2+]i imaging was done using a confocal microscope with a 40X water immersion objective lens with a numerical aperture of 0.8.
<ol>
	<li>Between 1 and 5 hr after slicing, place slice in the recording chamber and superfuse with oxygenated recording buffer at room temperature with flow rate of 1 - 2 ml per min. Then place a platinum harp with nylon strings on top of the slice to minimize movement during the experiment.</li>
	<li>Visualize and locate the dorsal striatum with bright field luminescence under a 10X water immersion objective.</li>
	<li>Switch to the 40X water immersion objective lens, and turn on the 488&#160;nm line of the Argon laser. By changing the focal plane, find cells located about 20 &#956;m below the slice surface, displaying basal fluorescence in soma, branches and branchlets. Of note, compromised cells usually display fluorescence level above average, which should be avoided.</li>
	<li>Use &#8216;frame scan&#8217; mode to start scanning. Usually, the laser intensity adjusted to 0.5 &#8211; 5% of the maximum output is sufficient to image [Ca2+]i with cyto-GCaMP3. For monitoring [Ca2+]i dynamics in the whole territory of astrocytes, the &#8216;frame scan&#8217; with a scan rate of 1&#160;sec per frame or faster is recommended, but this needs to be decided depending on the experiment in question.</li>
	<li>If necessary, to image [Ca2+]i in processes, use digital magnification of 2 - 3 fold to monitor 1 - 2 astrocytes in the whole field of view.</li>
	<li>To avoid saturated pixels during recording, use &#8216;HiLo&#8217; lookup mode to pseudocolor the cells. Always start with a &#8220;pilot recording&#8221; for about 5 min, to test if red color appears as indication of saturation. If so, lower the laser output power, or lower the PMT/Gain/offset values and perform another pilot recording to test the pixel values are in the non saturated range. Typically, the combined parameters used for imaging astrocyte [Ca2+]i in the striatum are as follows: laser output power 0.5 - 5% (of 10 mW), PMT 550 - 650 Volts, Gain 2.0 - 3.5X, and offset 0 - 5%.</li>
</ol>

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V., 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=Astrocytes+control+breathing+through+pH-dependent+release+of+ATP.">Astrocytes control breathing through pH-dependent release of ATP.</a> Science. 329, 571-575 (2010).</li><li>Florian, C., Vecsey, C. G., Halassa, M. M., Haydon, P. G., Abel, T. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Astrocyte-derived+adenosine+and+A1+receptor+activity+contribute+to+sleep+loss-induced+deficits+in+hippocampal+synaptic+plasticity+and+memory+in+mice.">Astrocyte-derived adenosine and A1 receptor activity contribute to sleep loss-induced deficits in hippocampal synaptic plasticity and memory in mice.</a> J Neurosci. 31, 6956-6962 (2011).</li><li>Shigetomi, E., Jackson-Weaver, O., Huckstepp, R. T., O'Dell, T. J., Khakh, B. 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D., Agulhon, C., Schmidt, E., Oheim, M., Ropert, N. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=New+tools+for+investigating+astrocyte-to-neuron+communication.">New tools for investigating astrocyte-to-neuron communication.</a> Frontiers in Cellular Neuroscience. 7, (2013).</li><li>Davila, D., Thibault, K., Fiacco, T. A., Agulhon, C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Recent+molecular+approaches+to+understanding+astrocyte+function+in+vivo.">Recent molecular approaches to understanding astrocyte function in vivo.</a> Front Cell Neurosci. 7, 272 (2013).</li><li>Paxinos, G., Franklin, K. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=.">.</a> The Mouse Brain in Stereotaxic Coordinates. , (2012).</li><li>Perea, G., Yang, A., Boyden, E. 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I., 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=Selective+induction+of+astrocytic+gliosis+generates+deficits+in+neuronal+inhibition.">Selective induction of astrocytic gliosis generates deficits in neuronal inhibition.</a> Nat Neurosci. 13, 584-591 (2010).</li><li>Zhang, Y., Barres, B. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Astrocyte+heterogeneity:+an+underappreciated+topic+in+neurobiology.">Astrocyte heterogeneity: an underappreciated topic in neurobiology.</a> Curr Opin Neurobiol. 20, 588-594 (2010).</li><li>Reimsnider, S., Manfredsson, F. P., Muzyczka, N., Mandel, R. 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The authors have nothing to disclose.

The methods described herein have allowed us to express cyto-GCaMP3 in striatal astrocytes in vivo for subsequent [Ca2+]i imaging in situ. This method has advantages over using transgenic or knock-in mice, including robust expression of the targeted protein, rapidity and flexibility of experimental implementation and anatomical specificity. The expression of GCaMP3 using AAV2/5 was found to be specific and robust. The combination of GFAP GfaABC1D promoter with AAV of the...

Astrocytes are ubiquitous and abundant glial cells of the brain. It is well established that astrocytes serve vital support and homeostatic roles including buffering of K+ concentration in the extracellular space, uptake of neurotransmitters as well as providing nutrients. However, recent studies show that they also display [Ca2+]i signals, which occur spontaneously and are increased by neuronal activity1. The existence of astrocyte [Ca2+]i signaling has been increasingly thought to trigger their communication with neurons, and as such has been interpreted as a form of &#8220;Ca2+ excitability&#8221; within astrocytes. The available data over the last two decades suggest two settings in which astrocytes and neurons may communicate, perhaps in a bidirectional manner. First, astrocytes often respond with an increase in [Ca2+]i when activated by neurotransmitters and neuromodulators released from neurons2. Second, [Ca2+]i increases within astrocytes cause the release of signaling molecules from astrocytes that in turn may affect neurons and blood vessels. Evidence suggests that molecules released from astrocytes lead to changes in the functions of synapses, circuits and ultimately behavior3-5 via astrocyte-to-neuron signaling. However, this remains a rapidly developing research area, and it has been argued that a better and detailed understanding of astrocyte [Ca2+]i is needed to resolve some of the current uncertainties6.
In past work, it was demonstrated that bulk loading of organic Ca2+ indicator dyes into astrocytes fails to reliably detect [Ca2+]i signals within entire astrocytes in culture and in situ7-10. These findings have been discussed by us and others6,11,12. The emerging picture is that [Ca2+]i signals within astrocyte processes (e.g., branches and branchlets), which are the primary sites for interactions with neurons and blood vessels, have rarely been explored in detail. Recently, the use of genetically encoded calcium indicators (GECIs) such as cytosolic GCaMP3, GCaMP5G and GCaMP6 and plasma membrane tethered versions (e.g., Lck-GCaMP3) has permitted the study of [Ca2+]i signals in small compartments of astrocytes such as thin processes, near the plasma membrane and within entire territories7,8. However, GECIs have one disadvantage over organic Ca2+ indicator dyes and that is the requirement for genetic methods to deliver the encoding genes selectively to astrocytes in vivo for periods of weeks for the GECIs to be appropriately expressed. Expression in vivo is typically achieved using transgenic mice, knock-in mice or with virus based delivery approaches. In the present JoVE article we report methods and procedures employed to deliver GECIs to striatal astrocytes using adeno-associated viruses. We focus on cyto-GCaMP3 as an example, but the same basic procedure works for any other GECI or fluorescent protein based reporter.

<table border="0" cellpadding="0" cellspacing="0" >
	<tbody>
		<tr >
			<td >Syringe Pump</td>
			<td>Harvard Apparatus</td>
			<td>704506</td>
			<td></td>
		</tr>
		<tr >
			<td >Glass Capillaries</td>
			<td >World Precision Instruments</td>
			<td >1B100-4</td>
			<td></td>
		</tr>
		<tr >
			<td >Micropipette puller</td>
			<td >Narishige</td>
			<td >PC-10</td>
			<td></td>
		</tr>
		<tr >
			<td >Micropipette grinder</td>
			<td >Narishige</td>
			<td >EG-40</td>
			<td></td>
		</tr>
		<tr >
			<td >pZac2.1 GfaABC1D.cyto-GCaMP3</td>
			<td >Addgene</td>
			<td>44331</td>
			<td >a plasmid sent to UPenn Vector Core for virus packaging</td>
		</tr>
		<tr >
			<td >I mL syringe</td>
			<td >BD</td>
			<td>309628</td>
			<td></td>
		</tr>
		<tr >
			<td >syringe needle</td>
			<td >BD</td>
			<td>305109</td>
			<td></td>
		</tr>
		<tr >
			<td >AAV2/5 virus</td>
			<td >UPenn vector core</td>
			<td >NA</td>
			<td></td>
		</tr>
		<tr >
			<td >Sudan red IV</td>
			<td >Sigma-Aldrich</td>
			<td >67386</td>
			<td></td>
		</tr>
		<tr >
			<td >Mineral oil</td>
			<td >CVS Pharmacy</td>
			<td>152355</td>
			<td></td>
		</tr>
		<tr >
			<td >Cryostat</td>
			<td >Leica</td>
			<td>CM3050 S</td>
			<td></td>
		</tr>
		<tr >
			<td >Stereotaxic instrument</td>
			<td>David Kopf Instruments</td>
			<td >900LS</td>
			<td></td>
		</tr>
		<tr >
			<td >High Speed Rotary Micromotor Kit</td>
			<td >FOREDOM</td>
			<td >K.1070</td>
			<td></td>
		</tr>
		<tr >
			<td >Paraformaldehyde</td>
			<td >Santa cruz biotechnology</td>
			<td >sc-281692</td>
			<td></td>
		</tr>
		<tr >
			<td >Super Glue</td>
			<td>Krazy&#174;Glue</td>
			<td>KG925</td>
			<td></td>
		</tr>
		<tr >
			<td >Microslicer</td>
			<td >Ted Pella</td>
			<td >DTK-Zero 1</td>
			<td></td>
		</tr>
		<tr >
			<td >Confocal microscopes</td>
			<td >Olympus</td>
			<td >FV300 and FV1000</td>
			<td></td>
		</tr>
		<tr >
			<td >Normal goat serum</td>
			<td >Vector</td>
			<td >S-1000</td>
			<td></td>
		</tr>
		<tr >
			<td >chicken anti-GFP</td>
			<td >Abcam</td>
			<td >ab13970</td>
			<td></td>
		</tr>
		<tr >
			<td >mouse anti-s100&#946;</td>
			<td >Sigma-Aldrich</td>
			<td >S2532</td>
			<td></td>
		</tr>
		<tr >
			<td >mouse anti-NeuN</td>
			<td >Millipore</td>
			<td >MAB377</td>
			<td></td>
		</tr>
		<tr >
			<td >mouse anti-glutamine synthetase</td>
			<td >Millipore</td>
			<td >MAB302</td>
			<td></td>
		</tr>
		<tr >
			<td >goat anti-mouse-Alexa546</td>
			<td >Invitrogen</td>
			<td >A11003</td>
			<td></td>
		</tr>
		<tr >
			<td >goat anti-chicken-Alexa488</td>
			<td >Invitrogen</td>
			<td >A11039</td>
			<td></td>
		</tr>
		<tr >
			<td >Microscope Slides</td>
			<td >Fisher Scientific</td>
			<td >12-550-15</td>
			<td></td>
		</tr>
		<tr >
			<td >Cover Glass</td>
			<td >Fisher Scientific</td>
			<td >12-548-5J</td>
			<td></td>
		</tr>
		<tr >
			<td >Mounting Medium</td>
			<td >Vector</td>
			<td >H-1000</td>
			<td></td>
		</tr>
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imaging intracellular ca2+ signals in striatal astrocytes from adult mice using genetically-encoded calcium indicators

Astrocytes display spontaneous intracellular Ca2+ concentration fluctuations ([Ca2+]i) and in several settings respond to neuronal excitation with enhanced [Ca2+]i signals. It has been proposed that astrocytes in turn regulate neurons and blood vessels through calcium-dependent mechanisms, such as the release of signaling molecules. However, [Ca2+]i imaging in entire astrocytes has only recently become feasible with genetically encoded calcium indicators (GECIs) such as the GCaMP series. The use of GECIs in astrocytes now provides opportunities to study astrocyte [Ca2+]i signals in detail within model microcircuits such as the striatum, which is the largest nucleus of the basal ganglia. In the present report, detailed surgical methods to express GECIs in astrocytes in vivo, and confocal imaging approaches to record [Ca2+]i signals in striatal astrocytes in situ, are described. We highlight precautions, necessary controls and tests to determine if GECI expression is selective for astrocytes and to evaluate signs of overt astrocyte reactivity. We also describe brain slice and imaging conditions in detail that permit reliable [Ca2+]i imaging in striatal astrocytes in situ. The use of these approaches revealed the entire territories of single striatal astrocytes and spontaneous [Ca2+]i signals within their somata, branches and branchlets. The further use and expansion of these approaches in the striatum will allow for the detailed study of astrocyte [Ca2+]i signals in the striatal microcircuitry.

For astrocyte specific expression of cyto-GCaMP3 in the striatum, we used adeno-associated virus (AAV) of the 5 serotype, and the GFAP GfaABC1D promoter (Figure 1A), which has been previously shown to drive robust GCaMP3 and reporter gene expression in hippocampal and cortical astrocytes8,14. Two weeks after virus microinjection into the mouse striatum, the mouse (~10 weeks old) was perfused and IHC was performed on thin brain sections to evaluate cyto-GCaMP3 expression in the stria...

Watch this Scientific Journal Video about Imaging Intracellular Ca2+ Signals in Striatal Astrocytes from Adult Mice Using Genetically-encoded Calcium Indicators at JoVE.com

Imaging Intracellular Ca2+ Signals in Striatal Astrocytes from Adult Mice Using Genetically-encoded Calcium Indicators

The properties and functions of astrocyte intracellular Ca2+ signals in the striatum remain incompletely explored. We describe methods to express genetically encoded calcium indicators in striatal astrocytes using adeno-associated viruses of serotype 2/5 (AAV2/5), as well as procedures to reliably image Ca2+ signals within striatal astrocytes in situ.

Imaging Intracellular Ca2+ Signals in Striatal Astrocytes from Adult Mice Using Genetically-encoded Calcium Indicators

Astrocytes display spontaneous intracellular Ca2+ concentration fluctuations ([Ca2+]i) and in several settings respond to neuronal excitation with ...

imaging-intracellular-ca2-signals-striatal-astrocytes-from-adult-mice

Research

JoVE Journal

Neuroscience

14.7K Views.  University of California Los Angeles. The properties and functions of astrocyte intracellular Ca2+ signals in the striatum remain incompletely explored. We describe methods to express genetically encoded calcium indicators in striatal astrocytes using adeno-associated viruses of serotype 2/5 (AAV2/5), as well as procedures to reliably image Ca2+ signals within striatal astrocytes in situ. 

Video: Imaging Intracellular Ca2+ Signals in Striatal Astrocytes from Adult Mice Using Genetically-encoded Calcium Indicators

Imaging Neuronal Responses in Slice Preparations of Vomeronasal Organ Expressing a Genetically Encoded Calcium Sensor

The vomeronasal organ (VNO) detects chemosensory signals that carry information about the social, sexual and reproductive status of the individuals within the same species 1,2. These intraspecies signals, the pheromones, as well as signals from some predators 3, activate the vomeronasal sensory neurons (VSNs) with high levels of specificity and sensitivity 4. At least three distinct families of G-protein coupled receptors, V1R, V2R and FPR 5-14, are expressed in VNO neurons to mediate the detection of the chemosensory cues. To understand how pheromone information is encoded by the VNO, it is critical to analyze the response profiles of individual VSNs to various stimuli and identify the specific receptors that mediate these responses.
The neuroepithelia of VNO are enclosed in a pair of vomer bones. The semi-blind tubular structure of VNO has one open end (the vomeronasal duct) connecting to the nasal cavity. VSNs extend their dendrites to the lumen part of the VNO, where the pheromone cues are in contact with the receptors expressed at the dendritic knobs. The cell bodies of the VSNs form pseudo-stratified layers with V1R and V2R expressed in the apical and basal layers respectively 6-8. Several techniques have been utilized to monitor responses of VSNs to sensory stimuli 4,12,15-19. Among these techniques, acute slice preparation offers several advantages. First, compared to dissociated VSNs 3,17, slice preparations maintain the neurons in their native morphology and the dendrites of the cells stay relatively intact. Second, the cell bodies of the VSNs are easily accessible in coronal slice of the VNO to allow electrophysiology studies and imaging experiments as compared to whole epithelium and whole-mount preparations 12,20. Third, this method can be combined with molecular cloning techniques to allow receptor identification.
Sensory stimulation elicits strong Ca2+ influx in VSNs that is indicative of receptor activation 4,21. We thus develop transgenic mice that express G-CaMP2 in the olfactory sensory neurons, including the VSNs 15,22. The sensitivity and the genetic nature of the probe greatly facilitate Ca2+ imaging experiments. This method has eliminated the dye loading process used in previous studies 4,21. We also employ a ligand delivery system that enables application of various stimuli to the VNO slices. The combination of the two techniques allows us to monitor multiple neurons simultaneously in response to large numbers of stimuli. Finally, we have established a semi-automated analysis pipeline to assist image processing.

The vomeronasal organ (VNO) detects intraspecies chemical signals that convey social and reproductive information. We have performed Ca2+ imaging experiments using transgenic mice expressing G-CaMP2 in VNO tissue. This approach allows us to analyze the complicated response patterns of the vomeronasal neurons to large numbers of pheromone stimuli.

The vomeronasal organ (VNO) detects chemosensory signals that carry information about the social, sexual and reproductive status of the individuals within ...

Simultaneous Recording of Calcium Signals from Identified Neurons and Feeding Behavior of Drosophila melanogaster

To study neuronal networks in terms of their function in behavior, we must analyze how neurons operate when each behavioral pattern is generated. Thus, simultaneous recordings of neuronal activity and behavior are essential to correlate brain activity to behavior. For such behavioral analyses, the fruit fly, Drosophila melanogaster, allows us to incorporate genetically encoded calcium indicators such as GCaMP1, to monitor neuronal activity, and to use sophisticated genetic manipulations for optogenetic or thermogenetic techniques to specifically activate identified neurons2-5. Use of a thermogenetic technique has led us to find critical neurons for feeding behavior (Flood et al., under revision). As a main part of feeding behavior, a Drosophila adult extends its proboscis for feeding6 (proboscis extension response; PER), responding to a sweet stimulus from sensory cells on its proboscis or tarsi. Combining the protocol for PER7 with a calcium imaging technique8 using GCaMP3.01, 9, I have established an experimental system, where we can monitor activity of neurons in the feeding center &#x2013; the suboesophageal ganglion (SOG), simultaneously with behavioral observation of the proboscis. I have designed an apparatus ("Fly brain Live Imaging and Electrophysiology Stage": "FLIES") to accommodate a Drosophila adult, allowing its proboscis to freely move while its brain is exposed to the bath for Ca2+ imaging through a water immersion lens. The FLIES is also appropriate for many types of live experiments on fly brains such as electrophysiological recording or time lapse imaging of synaptic morphology. Because the results from live imaging can be directly correlated with the simultaneous PER behavior, this methodology can provide an excellent experimental system to study information processing of neuronal networks, and how this cellular activity is coupled to plastic processes and memory.

Simultaneous Recording of Calcium Signals from Identified Neurons and Feeding Behavior of Drosophila melanogaster

The fruit fly, Drosophila melanogaster, extends its proboscis for feeding, responding to a sugar stimulus from its proboscis or tarsus. I have combined observations of the proboscis extension response (PER) with a calcium imaging technique, allowing us to monitor the activity of neurons in the brain, simultaneously with behavioral observation.

To study neuronal networks in terms of their function in behavior, we must analyze how neurons operate when each behavioral pattern is generated.  Thus, ...

Modeling Astrocytoma Pathogenesis In Vitro and In Vivo Using Cortical Astrocytes or Neural Stem Cells from Conditional, Genetically Engineered Mice

Current astrocytoma models are limited in their ability to define the roles of oncogenic mutations in specific brain cell types during disease pathogenesis and their utility for preclinical drug development. In order to design a better model system for these applications, phenotypically wild-type cortical astrocytes and neural stem cells (NSC) from conditional, genetically engineered mice (GEM) that harbor various combinations of floxed oncogenic alleles were harvested and grown in culture. Genetic recombination was induced in vitro using adenoviral Cre-mediated recombination, resulting in expression of mutated oncogenes and deletion of tumor suppressor genes. The phenotypic consequences of these mutations were defined by measuring proliferation, transformation, and drug response in vitro. Orthotopic allograft models, whereby transformed cells are stereotactically injected into the brains of immune-competent, syngeneic littermates, were developed to define the role of oncogenic mutations and cell type on tumorigenesis in vivo. Unlike most established human glioblastoma cell line xenografts, injection of transformed GEM-derived cortical astrocytes into the brains of immune-competent littermates produced astrocytomas, including the most aggressive subtype, glioblastoma, that recapitulated the histopathological hallmarks of human astrocytomas, including diffuse invasion of normal brain parenchyma. Bioluminescence imaging of orthotopic allografts from transformed astrocytes engineered to express luciferase was utilized to monitor in vivo tumor growth over time. Thus, astrocytoma models using astrocytes and NSC harvested from GEM with conditional oncogenic alleles provide an integrated system to study the genetics and cell biology of astrocytoma pathogenesis in vitro and in vivo and may be useful in preclinical drug development for these devastating diseases.

Modeling Astrocytoma Pathogenesis In Vitro and In Vivo Using Cortical Astrocytes or Neural Stem Cells from Conditional, Genetically Engineered Mice

Phenotypically wild-type astrocytes and neural stem cells harvested from mice engineered with floxed, conditional oncogenic alleles and transformed via viral Cre-mediated recombination can be used to model astrocytoma pathogenesis in vitro and in vivo by orthotopic injection of transformed cells into brains of syngeneic, immune-competent littermates.

Current astrocytoma models are limited in their ability to define the roles of oncogenic mutations in specific brain cell types during disease ...

Imaging Membrane Potential with Two Types of Genetically Encoded Fluorescent Voltage Sensors

Genetically encoded voltage indicators (GEVIs) have improved to the point where they are beginning to be useful for in vivo recordings. While the ultimate goal is to image neuronal activity in vivo, one must be able to image activity of a single cell to ensure successful in vivo preparations. This procedure will describe how to image membrane potential in a single cell to provide a foundation to eventually image in vivo. Here we describe methods for imaging GEVIs consisting of a voltage-sensing domain fused to either a single fluorescent protein (FP) or two fluorescent proteins capable of F&#246;rster resonance energy transfer (FRET) in vitro. Using an image splitter enables the projection of images created by two different wavelengths onto the same charge-coupled device (CCD) camera simultaneously. The image splitter positions a second filter cube in the light path. This second filter cube consists of a dichroic and two emission filters to separate the donor and acceptor fluorescent wavelengths depending on the FPs of the GEVI. This setup enables the simultaneous recording of both the acceptor and donor fluorescent partners while the membrane potential is manipulated via whole cell patch clamp configuration. When using a GEVI consisting of a single FP, the second filter cube can be removed allowing the mirrors in the image splitter to project a single image onto the CCD camera.

A method for imaging changes in membrane potential using genetically encoded voltage indicators is described.

Genetically encoded voltage indicators (GEVIs) have improved to the point where they are beginning to be useful for in vivo recordings. While the ultimate ...

Optical Control of a Neuronal Protein Using a Genetically Encoded Unnatural Amino Acid in Neurons

Photostimulation is a noninvasive way to control biological events with excellent spatial and temporal resolution. New methods are desired to photo-regulate endogenous proteins expressed in their native environment. Here, we present an approach to optically control the function of a neuronal protein directly in neurons using a genetically encoded unnatural amino acid (Uaa). By using an orthogonal tRNA/aminoacyl-tRNA synthetase pair to suppress the amber codon, a photo-reactive Uaa 4,5-dimethoxy-2-nitrobenzyl-cysteine (Cmn) is site-specifically incorporated in the pore of a neuronal protein Kir2.1, an inwardly rectifying potassium channel. The bulky Cmn physically blocks the channel pore, rendering Kir2.1 non-conducting. Light illumination instantaneously converts Cmn into a smaller natural amino acid Cys, activating Kir2.1 channel function. We express these photo-inducible inwardly rectifying potassium (PIRK) channels in rat hippocampal primary neurons, and demonstrate that light-activation of PIRK ceases the neuronal firing due to the outflux of K+ current through the activated Kir2.1 channels. Using in utero electroporation, we also express PIRK in the embryonic mouse neocortex in vivo, showing the light-activation of PIRK in neocortical neurons. Genetically encoding Uaa imposes no restrictions on target protein type or cellular location, and a family of photoreactive Uaas is available for modulating different natural amino acid residues. This technique thus has the potential to be generally applied to many neuronal proteins to achieve optical regulation of different processes in brains. The current protocol presents an accessible procedure for intricate Uaa incorporation in neurons in vitro and in vivo to achieve photo control of neuronal protein activity on the molecular level.

Here, a procedure to selectively activate a neuronal protein with a short pulse of light by genetically encoding a photo-reactive unnatural amino acid into a target neuronal protein expressed in neurons in culture or in vivo is presented.

Photostimulation is a noninvasive way to control biological events with excellent spatial and temporal resolution. New methods are desired to ...

Dissection of Local Ca2+ Signals in Cultured Cells by Membrane-targeted Ca2+ Indicators

Calcium ion (Ca2+) is a universal intracellular messenger molecule that drives multiple signaling pathways, leading to diverse biological outputs. The coordination of two Ca2+ signal sources&#8212;&#8220;Ca2+ influx&#8221; from outside the cell and &#8220;Ca2+ release&#8221; from the intracellular Ca2+ store endoplasmic reticulum (ER)&#8212;is considered to underlie the diverse spatio-temporal patterns of Ca2+ signals that cause multiple biological functions in cells. The purpose of this protocol is to describe a new Ca2+ imaging method that enables monitoring of the very moment of &#8220;Ca2+ influx&#8221; and &#8220;Ca2+ release&#8221;. OER-GCaMP6f is a genetically encoded Ca2+ indicator (GECI) comprising GCaMP6f, which is targeted to the ER outer membrane. OER-GCaMP6f can monitor Ca2+ release at a higher temporal resolution than conventional GCaMP6f. Combined with plasma membrane-targeted GECIs, the spatio-temporal Ca2+ signal pattern can be described at a subcellular resolution. The subcellular-targeted Ca2+ indicators described here are, in principle, available for all cell types, even for the in vivo imaging of Caenorhabditis elegans neurons. In this protocol, we introduce Ca2+ imaging in cells from cell lines, neurons, and glial cells in dissociated primary cultures, and describe the preparation of frozen stock of rat cortical neurons.

Dissection of Local Ca2+ Signals in Cultured Cells by Membrane-targeted Ca2+ Indicators

Here we present a protocol for Ca2+ imaging in neurons and glial cells, which enables the dissection of Ca2+ signals at subcellular resolution. This process is applicable to all cell types that allow the expression of genetically encoded Ca2+ indicators.

Calcium ion (Ca2+) is a universal intracellular messenger molecule that drives multiple signaling pathways, leading to diverse biological outputs. The ...

Single Synapse Indicators of Glutamate Release and Uptake in Acute Brain Slices from Normal and Huntington Mice

Synapses are highly compartmentalized functional units that operate independently on each other. In Huntington&#39;s disease (HD) and other neurodegenerative disorders, this independence might be compromised due to insufficient glutamate clearance and the resulting spill-in and spill-out effects. Altered astrocytic coverage of the presynaptic terminals and/or dendritic spines as well as a reduced size of glutamate transporter clusters at glutamate release sites have been implicated in the pathogenesis of diseases resulting in symptoms of dys-/hyperkinesia. However, the mechanisms leading to the dysfunction of glutamatergic synapses in HD are not well understood. Improving and applying synapse imaging we have obtained data shedding new light on the mechanisms impeding the initiation of movements. Here, we describe the principle elements of a relatively inexpensive approach to achieve single synapse resolution by using the new genetically encoded ultrafast glutamate sensor iGluu, wide-field optics, a scientific CMOS (sCMOS) camera, a 473 nm laser and a laser positioning system to evaluate the state of corticostriatal synapses in acute slices from age appropriate healthy or diseased mice. Glutamate transients were constructed from single or multiple pixels to obtain estimates of i) glutamate release based on the maximal elevation of the glutamate concentration [Glu] next to the active zone and ii) glutamate uptake as reflected in the time constant of decay (TauD) of the perisynaptic [Glu]. Differences in the resting bouton size and contrasting patterns of short-term plasticity served as criteria for the identification of corticostriatal terminals as belonging to the intratelencephalic (IT) or the pyramidal tract (PT) pathway. Using these methods, we discovered that in symptomatic HD mice ~40% of PT-type corticostriatal synapses exhibited insufficient glutamate clearance, suggesting that these synapses might be at risk to excitotoxic damage. The results underline the usefulness of TauD as a biomarker of dysfunctional synapses in Huntington mice with a hypokinetic phenotype.

We present a protocol to evaluate the balance between glutamate release and clearance at single corticostriatal glutamatergic synapses in acute slices from adult mice. This protocol uses the fluorescent sensor iGluu for glutamate detection, a sCMOS camera for signal acquisition and a device for focal laser illumination.

Synapses are highly compartmentalized functional units that operate independently on each other. In Huntington&#39;s disease (HD) and other ...

Visualizing and Analyzing Intracellular Transport of Organelles and Other Cargos in Astrocytes

Astrocytes are among the most abundant cell types in the adult brain, where they play key roles in a multiplicity of functions. As a central player in brain homeostasis, astrocytes supply neurons with vital metabolites and buffer extracellular water, ions, and glutamate. An integral component of the &#8220;tri-partite&#8221; synapse, astrocytes are also critical in the formation, pruning, maintenance, and modulation of synapses. To enable these highly interactive functions, astrocytes communicate among themselves and with other glial cells, neurons, the brain vasculature, and the extracellular environment through a multitude of specialized membrane proteins that include cell adhesion molecules, aquaporins, ion channels, neurotransmitter transporters, and gap junction molecules. To support this dynamic flux, astrocytes, like neurons, rely on tightly coordinated and efficient intracellular transport. Unlike neurons,&#160;where intracellular trafficking has been extensively delineated, microtubule-based transport in astrocytes has been less studied. Nonetheless, exo- and endocytic trafficking of cell membrane proteins and intracellular organelle transport orchestrates astrocytes&#8217; normal biology, and these processes are often affected in disease or in response to injury. Here we present a straightforward protocol to culture high quality murine astrocytes, to fluorescently label astrocytic proteins and organelles of interest, and to record their intracellular transport dynamics using time-lapse confocal microscopy. We also demonstrate how to extract and quantify relevant transport parameters from the acquired movies using available image analysis software (i.e., ImageJ/FIJI) plugins.

Here we describe an in vitro live-imaging method to visualize intracellular transport of organelles and trafficking of plasma membrane proteins in murine astrocytes. This protocol also presents an image analysis methodology to determine cargo transport itineraries and kinetics.

Astrocytes are among the most abundant cell types in the adult brain, where they play key roles in a multiplicity of functions. As a central player in ...

Minimizing Hypoxia in Hippocampal Slices from Adult and Aging Mice

Acute hippocampal slices have enabled generations of neuroscientists to explore synaptic, neuronal, and circuit properties in detail and with high fidelity. Exploration of LTP and LTD mechanisms, single neuron dendritic computation, and experience-dependent changes in circuitry, would not have been possible without this classical preparation. However, with a few exceptions, most basic research using acute hippocampal slices has been performed using slices from rodents of relatively young ages, ~P20-P40, even though synaptic and intrinsic excitability mechanisms have a long developmental tail that reaches past P60. The main appeal of using young hippocampal slices is preservation of neuronal health aided by higher tolerance to hypoxic damage. However, there is a need to understand neuronal function at more mature stages of development, further accentuated by the development of various animal models of neurodegenerative diseases that require an aging brain preparation. Here we describe a modification to an acute hippocampal slice preparation that reliably delivers healthy slices from adult and aging mouse hippocampi. The protocol&#8217;s critical steps are transcardial perfusion and cutting with ice-cold sodium-free NMDG-aSCF. Together, these steps attenuate the hypoxia-induced drop in ATP upon decapitation, as well as cytotoxic edema caused by passive sodium fluxes. We demonstrate how to cut transversal slices of hippocampus plus cortex using a vibrating microtome. Acute hippocampal slices obtained in this way are reliably healthy over many hours of recording, and are appropriate for both field-recordings and targeted patch-clamp recordings, including targeting of fluorescently labeled neurons.

This is a protocol for acute slice preparation from adult and aging mouse hippocampi that takes advantage of transcardial perfusion and slice cutting with ice-cold NMDG-aCSF to reduce hypoxic damage to the tissue. The resulting slices stay healthy over many hours, and are suitable for long-term patch-clamp and field-recordings.

Acute hippocampal slices have enabled generations of neuroscientists to explore synaptic, neuronal, and circuit properties in detail and with high ...

In vivo Calcium Imaging in Mouse Inferior Olive

Inferior olive (IO), a nucleus in the ventral medulla, is the only source of climbing fibers that form one of the two input pathways entering the cerebellum. IO has long been proposed to be crucial for motor control and its activity is currently considered to be at the center of many hypotheses of both motor and cognitive functions of the cerebellum. While its physiology and function have been relatively well studied on single-cell level in vitro, presently there are no reports on the organization of the IO network activity in living animals. This is largely due to the extremely challenging anatomical location of the IO, making it difficult to subject to conventional fluorescent imaging methods, where an optic path must be created through the entire brain located dorsally to the region of interest.
Here we describe an alternative method for obtaining state-of-the-art -level calcium imaging data from the IO network. The method takes advantage of the extreme ventral location of the IO and involves a surgical procedure for inserting a gradient-refractive index (GRIN) lens through the neck viscera to come into contact with the ventral surface of the calcium sensor GCaMP6s-expressing IO in anesthetized mice. A representative calcium imaging recording is shown to demonstrate the feasibility to record IO neuron activity after the surgery. While this is a non-survival surgery and the recordings must be conducted under anesthesia, it avoids damage to life-critical brainstem nuclei and allows conducting large variety of experiments investigating spatiotemporal activity patterns and input integration in the IO. This procedure with modifications could be used for recordings in other, adjacent regions of the ventral brainstem.

In vivo Calcium Imaging in Mouse Inferior Olive

We present a protocol to expose the brainstem of adult mouse from the ventral side. By using a gradient-refractive index lens with a miniature microscope, calcium imaging can be used to examine the activity of inferior olive neural somata in vivo.