This work was partially supported by National Institute of Deafness and Other Communications Disorders Awards R03-DC-012125 to D. A. Llano and F31-DC-013501 to B. J. Slater as well as the Carver Foundation.
The authors would like to thank Jason MacLean and Matthew Banks for technical advice with calcium imaging.

All procedures were approved by the Institutional Animal Care and Use Committee at the University of Illinois. All animals were housed in animal care facilities approved by the American Association for Accreditation of Laboratory Animal Care. Every attempt was made to minimize the number of animals used and to reduce suffering at all stages of the study.
1. Preparation for and Removal of Brain from Mouse for Slicing
<ol>
	<li>Prepare for perfusion and slice incubation.
	<ol>
		<li>Approximately 30 min before slicing, prepare high sucrose cutting solution and low calcium aCSF for incubation of the slice prior to recording or imaging.</li>
		<li>Lay out all necessary tools (large blunt end scissors, small spring scissors, anatomic forceps, large scissors or guillotine, iris scissors, jewelers forceps, 10 ml&#160;syringe with 27G&#160;x 1/2 inch needle, small piece of 1 cm x 2 cm filter paper, and a broken tipped Pasteur pipette for slice transfer) for perfusion and slicing, and prepare perfusion tray.</li>
		<li>Set up incubating chamber with oxygenated aCSF in a 32 oC water bath and a culture dish filled with cutting solution.</li>
	</ol>
	</li>
	<li>Prepare cutting stage for slicing.
	<ol>
		<li>Cut a small piece of 3% agar approximately 1 cm3 to use as a backstop for the brain and 1.5 cm x 1.5 mm x 3 mm for a bump to be used to prop up the IC.</li>
		<li>Glue the backstop onto the stage with cyanoacrylate adhesive as well as the bump on the right side of the backstop such that they form an 80o angle.</li>
	</ol>
	</li>
	<li>Remove the brain.
	<ol>
		<li>Deeply anesthetize a p12-p20 (ideally p14-18) mouse with ketamine 100 mg/kg and xylaxine 3 mg/kg (or comparable ethics committee approved procedure). 
		Note: Confirm proper anesthesia level via lack of response to toe pinch. As the animal is under anesthesia briefly, ophthalmic ointment is unnecessary.</li>
		<li>Using blunt end scissors, expose the ribcage from the xiphoid process to the neck.
		<ol>
			<li>Find the xiphoid process, and make a horizontal cut of approximately 1.5 cm.</li>
			<li>Make two vertical cuts from the ends of the previous horizontal cut to the shoulders, approximately 1.5 cm each.</li>
		</ol>
		</li>
		<li>Cut through the diaphragm and the costochondral junctions to expose the heart.</li>
		<li>With small spring scissors, make a small, 2-5 mm cut in the right atrium, from the ventral to dorsal side of the heart.</li>
		<li>Using a 10 ml&#160;syringe with a 27G&#160;x 1/2 inch needle, inject the left ventricle and quickly perfuse the animal with high sucrose cutting solution.</li>
		<li>Once the blood runs clear, use larger scissors to remove the head. 
		Note: We have not systematically assessed the utility of perfusion. However, in our experience, transcardiac perfusion in mice this young can be done with nearly 100% success, and does have the benefit of eliminating red blood cells, which can fluoresce and interfere with imaging.</li>
		<li>Cut the skin down the midline to expose the skull.</li>
		<li>Using bent iris scissors, cut the skull between the eyes, then starting from the cut between the eyes, carefully cut from anterior to posterior along the midline suture taking care not to damage the brain underneath. 
		Note: The dura usually comes off with the skull. If necessary, remove the dura before carefully removing the brain.</li>
		<li>Using jewelers forceps, pry open the skull and carefully remove the brain, then place the brain in cutting solution. Take care to avoid damaging the cortices.</li>
	</ol>
	</li>
</ol>
2. Preparing Brain for Slicing
<ol>
	<li>Preparing brain for mounting on vibratome stage.
	<ol>
		<li>Using a slide marked with two lines at 90o and a diagonal line at 17o from the top left to bottom right, intersecting where the two lines meet, place the brain dorsal side up, using a razor blade, remove 2-4 mm of the rostral end brain creating a flat surface.</li>
		<li>Place the brain caudal side up on the newly created flat surface, and align the dorsal surface of the brain with the horizontal and the midline of the brain with the vertical line.</li>
		<li>Align the razor blade with the 17o line, tilt the razor at a 30o angle and remove approximately 3 mm of the right cortex in a double diagonal cut (17o from horizontal plane and 60o from the coronal).</li>
	</ol>
	</li>
	<li>Mounting brain on vibratome stage.
	<ol>
		<li>Place a small piece of filter paper 1 cm x 2 cm on the ventral side of the brain so that the long dimension is perpendicular to the midline.</li>
		<li>Carefully apply a small amount of cyanoacrylate adhesive to the area in front of the backstop (and to the left of the bump).</li>
		<li>Place the double diagonally-cut brain face on to the glue so that the caudal part of the brain and hindbrain are propped upon the bump and the right side of the brain is against the backstop.</li>
		<li>Delicately press down on the brain, ensuring that the entire surface is in contact with the slicing chamber.</li>
	</ol>
	</li>
</ol>
3. Obtaining the Colliculo-thalamocortical Slice
<ol>
	<li>Quickly take the cutting stage and place in the vibratome, fill the stage with cutting solution.</li>
	<li>Align the blade with the top (ventral side) of the brain.</li>
	<li>Remove 1-1.5 mm from the top of the brain, remove 300-500 &#181;m slices and assess depth after removal. 
	Note: When the IC, the MGB, and the lateral geniculate nucleus (LGN) are all visible, and the dentate gyrus forms a &#39;C&#39; shape take one to two 600 &#181;m slices. See Figure 3A.</li>
	<li>Place slices in heated (32 oC) holding chamber.</li>
</ol>
4. Imaging of the Slice
<ol>
	<li>Preparation for flavoprotein autofluorescence imaging.
	<ol>
		<li>Prepare aCSF for perfusion of slice in a standard electrophysiological recording chamber, bubble aCSF with 95% O2/5% CO2.</li>
		<li>Pull glass electrode for stimulation. Note that multiple different stimulating electrodes and configurations (glass, tungsten, carbon fiber, monopolar, bipolar) all work very well in conjunction with flavoprotein autofluorescence imaging.</li>
		<li>Turn on computer, camera, micromanipulator, light source, stimulation software, image capture software.</li>
		<li>Perfuse the recording chamber with oxygenated aCSF, place custom bridge (design available in Supplemental Materials) to elevate slice in chamber. 
		Note: Perfusion rate can vary with any individual set up; 5-15 ml/min is used here.</li>
		<li>Place slice in chamber, lower the liquid level in the chamber to prevent the slice from lifting off the bridge while placing specially constructed harp over slice, taking care to avoid placing harp strings over pathways of interest (Figure 1A). Increase liquid level to approximately 1-2 mm above slice to maintain aCSF flow above and below slice. 
		Note: If repositioning is necessary, use the broken end Pasteur pipette or allow the aCSF level to rise and use forceps with extreme care.</li>
		<li>Insert silver chloride electrode into glass electrode filled with aCSF, insert the glass electrode into micromanipulator and connect to stimulus isolator, and carefully place glass electrode in IC/white matter leading to thalamus(see Figure 1A).</li>
	</ol>
	</li>
	<li>Flavoprotein autofluorescence image acquisition.
	<ol>
		<li>Collect images at 4 Hz (using an infinity-corrected 2X macro objective (NA 0.13)) and a camera for 105 sec&#160;while electrically stimulating (each stimulation consists of 1 sec&#160;of 40 Hz 2 msec pulses tissue at 0.05 Hz five times) while illuminating the tissue with blue light 470-490 nm and capturing above 515 nm. Ensure that the image is neither too dark, nor blown out see Figure 3 left column for reference. 
		Note: Collection time, collection frequency, and stimulation frequency can be changed to suit the experiment, the custom program included in supplementary materials can be modified as such.</li>
		<li>Export images to Matlab, and using the custom written program available in supplementary materials to analyze spectral power of the images at 0.05 Hz. The resulting image will show connected pathways. 
		Note: The custom program uses a fast Fourier transform of the pixel values in the time series to produce the image.</li>
	</ol>
	</li>
	<li>Preparation for calcium imaging.
	<ol>
		<li>Prepare small incubation chamber using a 3 cm culture dish and raised culture membrane to allow aCSF to reach both the top and bottom of the slice, and place on a heating pad to maintain temperature at approximately 32 oC.</li>
		<li>Fill small incubation chamber with warm aCSF and slowly bubble with 95% O2/5% CO2.</li>
		<li>Mix 2 &#181;l&#160;of pluronic F-127 acid and 50 &#181;g of Fura-2AM dissolved in 48 &#181;l&#160;of DMSO.</li>
		<li>Place slice in small incubation chamber on raised membrane and carefully using a micropipette add staining mixture directly above the MGB (or other structure of interest).</li>
		<li>Cover slices to prevent bleaching prior to imaging, and allow slices to incubate for 45 min - 1 hr.</li>
		<li>Remove slices to heated holding chamber for 10-15 min&#160;to wash off excess material.</li>
		<li>Follow steps 4.1.1 to 4.1.6 for preparation to stimulate and image the colliculo-thalamocortical slice.</li>
	</ol>
	</li>
	<li>Calcium image acquisition.
	<ol>
		<li>Collect images at 10 Hz (using an 20X water immersion objective) for 25 sec&#160;while electrically stimulating (each stimulation consists of one 2 msec pulse) tissue at 0.2 Hz five times while illuminating the tissue with 365 nm light and capturing fluorescence above 510 nm.</li>
		<li>Export images to Matlab and using the custom written program available in supplementary materials to analyze spectral power of the images at 0.2 Hz. The resulting image will show active cells.</li>
	</ol>
	</li>
</ol>

<ol>
	<li>Llano, D. A., Slater, B. J., Lesicko, A. M., Stebbings, K. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=An+auditory+colliculothalamocortical+brain+slice+preparation+in+mouse.">An auditory colliculothalamocortical brain slice preparation in mouse.</a> J. Neurosci. 111, 197-207 (2014).</li><li>Agmon, A., Connors, B. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Thalamocortical+responses+of+mouse+somatosensory+(barrel)+cortex+in+vitro.">Thalamocortical responses of mouse somatosensory (barrel) cortex in vitro.</a> Neuro. 41, 365-379 (1991).</li><li>Cruikshank, S. J., Rose, H. J., Metherate, R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Auditory+Thalamocortical+Synaptic+Transmission+In+Vitro.">Auditory Thalamocortical Synaptic Transmission In Vitro.</a> J Neurophysiol. 87, 361-384 (2002).</li><li>Lee, C. C., Sherman, S. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Topography+and+physiology+of+ascending+streams+in+the+auditory+tectothalamic+pathway.">Topography and physiology of ascending streams in the auditory tectothalamic pathway.</a> PNAS. 107, 372-377 (2010).</li><li>MacLean, J. N., Fenstermaker, V., Watson, B. O., Yuste, R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+visual+thalamocortical+slice.">A visual thalamocortical slice.</a> Nat meth. 3, 129-134 (2006).</li><li>Crabtree, J. W., Isaac, J. T. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=New+intrathalamic+pathways+allowing+modality-related+and+cross-modality+switching+in+the+dorsal+thalamus.">New intrathalamic pathways allowing modality-related and cross-modality switching in the dorsal thalamus.</a> J. Neurosci. 22, 8754-8761 (2002).</li><li>McAlonan, K., Cavanaugh, J., Wurtz, R. H. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Attentional+Modulation+of+Thalamic+Reticular+Neurons.">Attentional Modulation of Thalamic Reticular Neurons.</a> J. Neurosci. 26, 4444-4450 (2006).</li><li>Weese, G. D., Phillips, J. M., Brown, V. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Attentional+Orienting+Is+Impaired+by+Unilateral+Lesions+of+the+Thalamic+Reticular+Nucleus+in+the+Rat.">Attentional Orienting Is Impaired by Unilateral Lesions of the Thalamic Reticular Nucleus in the Rat.</a> J. Neurosci. 19, 10135-10139 (1999).</li><li>Zikopoulos, B., Barbas, H. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Prefrontal+Projections+to+the+Thalamic+Reticular+Nucleus+form+a+Unique+Circuit+for+Attentional+Mechanisms.">Prefrontal Projections to the Thalamic Reticular Nucleus form a Unique Circuit for Attentional Mechanisms.</a> J. Neurosci. 26, 7348-7361 (2006).</li><li>Kalatsky, V. A., Stryker, M. P. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=New+Paradigm+for+Optical+Imaging:+Temporally+Encoded+Maps+of+Intrinsic+Signal.">New Paradigm for Optical Imaging: Temporally Encoded Maps of Intrinsic Signal.</a> Neuron. 38, 529-545 (2003).</li><li>Llano, D. A., Turner, J., Caspary, D. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Diminished+cortical+inhibition+in+an+aging+mouse+model+of+chronic+tinnitus.">Diminished cortical inhibition in an aging mouse model of chronic tinnitus.</a> J. Neurosci. 32, 16141-16148 (2012).</li><li>Ehret, G. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Behavioural+studies+on+auditory+development+in+mammals+in+relation+to+higher+nervous+system+functioning.">Behavioural studies on auditory development in mammals in relation to higher nervous system functioning.</a> Acta otolaryngol. 99, 31-40 (1985).</li><li>Mikaelian, D., Ruben, R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Development+of+hearing+in+the+normal+CBA-J+mouse:+correlation+of+physiological+observations+with+behavioral+responses+and+with+cochlear+anatomy.">Development of hearing in the normal CBA-J mouse: correlation of physiological observations with behavioral responses and with cochlear anatomy.</a> Acta. 59, 451-461 (1965).</li></ol>

The authors have nothing to disclose.

This protocol describes improvements upon a previously described colliculo-thalamocortical brain slice in p12-20 mouse to study information flow in the auditory system1. This method has a number of advantages over other, similar, brain slice preparations by retaining connections between more brain areas in a single slice, which gives investigators new tools to understand the interaction and interplay between auditory nuclei in the forebrain. There have been a few key modifications in this protocol, compared to...

In the auditory system, although there is substantial processing of information between the sensory periphery and the inferior colliculus, there is considerable additional processing before it reaches the auditory cortex. We know very little about how that processing is done and therefore little about how that transformation allows the brain to interpret incoming sensory information. With the exception of olfaction, each of the senses has a very similar organization with peripheral signals initially being relayed with high fidelity which declines as the signal ascends to the cortex. The cortex then sends projections to the lower structures to further modulate the incoming information. This complex system has been studied in a variety of ways in vivo as well as in a number of in vitro preparations. In the former, all connections are intact, enabling the researcher to probe any set of connections, while controlling the sensory input and measuring output in any given area. With this approach, there is little to no control of the large variety of other inputs, including other sensory inputs, arousal, and attention, giving rise to an intensely complex output. In vitro, brain slices have been cut to capture either a single set of projections, or two connected brain areas, which allow researchers to stimulate and evaluate various afferents or brain areas. These are often either thalamocortical or tectothalamic slices where either the input to the thalamus or the thalamus and its output to the cortex are preserved2-5. These preparations allow for a wide variety of pharmacological, electrical, and optogenetic manipulations. However with only two brain regions, they primarily evaluate the transfer of information and lack the ability to evaluate the transformation of information as it passes through the thalamus. Also the reticulo-thalamic projection, which may play a role in attention modulation6-9 is present in this slice. Here we demonstrate improvements upon our previous preparation1, which allows the investigator control of various inputs to the thalamus to give a unique perspective of how the thalamus gates and filters information. We couple this novel slice preparation with flavoprotein autofluorescence imaging for assessing slice connectivity and large-scale activation analysis, calcium imaging in the thalamus for neuronal population analysis, and single cell recording to measure the impact of the various inputs on a single cell level.
To assist in maintaining these widespread connections we have also developed a number of modifications of the normal slice anchor (a.k.a. &#8220;harp&#8221;) for holding the brain slice in place and a bridge to elevate the slice for enhanced perfusion. The harp is designed in a modified horseshoe shape to surround the slice and allow for customizable attachment points for the harp strings. Three strings are attached such that i) one lies horizontally along the medial edge of the slice, ii) one extends from the caudal edge of the IC to the caudal edge of the AC and iii) one extends diagonally from the medial edge of the slice to an area rostral to the AC (see Figure 1A). Small indentations in the frame for gluing (with cyanoacrylate glue) of the harp strings allow for a decreased amount of pressure on the slice to help maintain slice integrity (see Figure 1B). By using three dimensional printing, we are able to custom design harps to our unique specifications, as well as bridges which allow for ideal flow of artificial cerebrospinal fluid (aCSF) above and below the tissue. This also maintains large areas for light to penetrate the tissue for patch clamp electrophysiology.

<table border="0" cellpadding="0" cellspacing="0">
	<tbody>
		<tr>
			<td>High sucrose cutting solution</td>
			<td></td>
			<td></td>
			<td>in mM: 206 sucrose, 10.0 MgCl2, 11.0 glucose, 1.25 NaH2PO4, 26 NaHCO3, 0.5 CaCl2, 2.5 KCl, pH 7.4</td>
		</tr>
		<tr>
			<td>Low calcium aCSF</td>
			<td></td>
			<td></td>
			<td>in mM: 126 NaCl, 3.0 MgCl2, 10.0 glucose, 1.25 NaH2PO4, 26 NaHCO3, 1.0 CaCl2, 2.5 KCl, pH 7.4</td>
		</tr>
		<tr>
			<td>aCSF</td>
			<td></td>
			<td></td>
			<td>in mM: 126 NaCl, 2.0 MgCl2, 10.0 glucose, 1.25 NaH2PO4, 26 NaHCO3, 2.0 CaCl2, 2.5 KCl, pH 7.4</td>
		</tr>
		<tr>
			<td>Stimulus Isolator</td>
			<td>World Precision Instruments</td>
			<td>A360</td>
			<td></td>
		</tr>
		<tr>
			<td>DMSO</td>
			<td>Life Technologies</td>
			<td>D12345</td>
			<td>Lot: 1572C502</td>
		</tr>
		<tr>
			<td>Fura-2AM</td>
			<td>Life Technologies</td>
			<td>F1201</td>
			<td>Lot: 144912</td>
		</tr>
		<tr>
			<td>Pluronic F-127</td>
			<td>Life Technologies</td>
			<td>P3000MP</td>
			<td>Lot: 1499369</td>
		</tr>
		<tr>
			<td>Large culture dish</td>
			<td>Fisherbrand</td>
			<td>08-757-13</td>
			<td>100x15mm culture dish</td>
		</tr>
		<tr>
			<td>Small culture dish</td>
			<td>Falcon</td>
			<td align="right">353001</td>
			<td>35x10mm culture dish</td>
		</tr>
		<tr>
			<td>Raised culture membrane</td>
			<td>Millicell</td>
			<td>PICMORG50</td>
			<td>Used to maintain oxygenated fluid perfusion on both sides of slice.</td>
		</tr>
		<tr>
			<td>Flavoprotein imaging fluorescence cube</td>
			<td>Olympus</td>
			<td>UMNIB</td>
			<td>470&#8211;490 nm excitation, 505 nm dichroic, 515 nm emission long pass.&#160; We have found that virtually any green fluorescence protein filter cube will work here.</td>
		</tr>
		<tr>
			<td>Calcium imaging fluorescence cube</td>
			<td>Omega Optical</td>
			<td>BX-18</td>
			<td>XF1005 365nm exitation, XF2001 400nm dichroic, XF3080 510nm emission</td>
		</tr>
		<tr>
			<td>Agar for blocking brain</td>
			<td></td>
			<td></td>
			<td>3% by weight in water</td>
		</tr>
		<tr>
			<td>Viper si Stereo Lithography Apparatus</td>
			<td>3D Systems</td>
			<td></td>
			<td></td>
		</tr>
	</tbody>
</table>

modification of a colliculo-thalamocortical mouse brain slice, incorporating 3-d printing of chamber components and multi-scale optical imaging

The ability of the brain to process sensory information relies on both ascending and descending sets of projections. Until recently, the only way to study these two systems and how they interact has been with the use of in vivo preparations. Major advances have been made with acute brain slices containing the thalamocortical and cortico-thalamic pathways in the somatosensory, visual, and auditory systems. With key refinements to our recent modification of the auditory thalamocortical slice1, we are able to more reliably capture the projections between most of the major auditory midbrain and forebrain structures: the inferior colliculus (IC), medial geniculate body (MGB), thalamic reticular nucleus (TRN), and the auditory cortex (AC). With portions of all these connections retained, we are able to answer detailed questions that complement the questions that can be answered with in vivo preparations. The use of flavoprotein autofluorescence imaging enables us to rapidly assess connectivity in any given slice and guide the ensuing experiment. Using this slice in conjunction with recording and imaging techniques, we are now better equipped to understand how information processing occurs at each point in the auditory forebrain as information ascends to the cortex, and the impact of descending cortical modulation. 3-D printing to build slice chamber components permits double-sided perfusion and broad access to networks within the slice and maintains the widespread connections key to fully utilizing this preparation.

An example of colliculo-thalamocortical mouse brain slice obtained in P15 mouse is shown in figure 2. The ideal slice will contain the four major midbrain and forebrain auditory structures IC, MGB, TRN, and AC, which are all activated when the IC is stimulated (Figure 2A). Using Fourier analysis, the spectral power is measured at the electrical stimulation frequency, with connected brain regions showing activity that is periodic and entrained at the stimulation frequency10. Th...

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Modification of a Colliculo-thalamocortical Mouse Brain Slice, Incorporating 3-D printing of Chamber Components and Multi-scale Optical Imaging

Using multiple angles to cut the mouse pup brain, we improve upon a previously-described acute brain slice which captures the connections between most of the major auditory midbrain and forebrain structures.

The ability of the brain to process sensory information relies on both ascending and descending sets of projections. Until recently, the only way to study ...

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Neuroscience

8.2K visualizações.  University of Illinois Urbana-Champaign.  O uso de vários ângulos para cortar o cérebro do rato filhote, podemos melhorar em cima de uma fatia cerebral aguda anteriormente descrito que captura as conexões entre a maioria dos principais mesencéfalo e prosencéfalo estruturas auditivas. 

Vídeo: Modification of a Colliculo-thalamocortical Mouse Brain Slice, Incorporating 3-D printing of Chamber Components and Multi-scale Optical Imaging

Scalable Fluidic Injector Arrays for Viral Targeting of Intact 3-D Brain Circuits

Our understanding of neural circuits--how they mediate the computations that subserve sensation, thought, emotion, and action, and how they are corrupted in neurological and psychiatric disorders--would be greatly facilitated by a technology for rapidly targeting genes to complex 3-dimensional neural circuits, enabling fast creation of "circuit-level transgenics." We have recently developed methods in which viruses encoding for light-sensitive proteins can sensitize specific cell types to millisecond-timescale activation and silencing in the intact brain. We here present the design and implementation of an injector array capable of delivering viruses (or other fluids) to dozens of defined points within the 3-dimensional structure of the brain (Figure. 1A, 1B). The injector array comprises one or more displacement pumps that each drive a set of syringes, each of which feeds into a polyimide/fused-silica capillary via a high-pressure-tolerant connector. The capillaries are sized, and then inserted into, desired locations specified by custom-milling a stereotactic positioning board, thus allowing viruses or other reagents to be delivered to the desired set of brain regions. To use the device, the surgeon first fills the fluidic subsystem entirely with oil, backfills the capillaries with the virus, inserts the device into the brain, and infuses reagents slowly (&lt;0.1 microliters/min). The parallel nature of the injector array facilitates rapid, accurate, and robust labeling of entire neural circuits with viral payloads such as optical sensitizers to enable light-activation and silencing of defined brain circuits. Along with other technologies, such as optical fiber arrays for light delivery to desired sets of brain regions, we hope to create a toolbox that enables the systematic probing of causal neural functions in the intact brain. This technology may not only open up such systematic approaches to circuit-focused neuroscience in mammals, and facilitate labeling of brain regions in large animals such as non-human primates, but may also open up a clinical translational path for cell-specific optical control prosthetics, whose precision may enable improved treatment of intractable brain disorders. Finally, such devices as described here may facilitate precisely-timed fluidic delivery of other payloads, such as stem cells and pharmacological agents, to 3-dimensional structures, in an easily user-customizable fashion.

Controlling and analyzing neural circuits in vivo would be facilitated by a technology for delivery of viruses and other reagents to desired 3-dimensional sets of brain regions. We demonstrate customized fluidic injector array fabrication, and delivery of virally-encoded optical sensitizers, enabling optical manipulation of complex brain circuits.

Our understanding of neural circuits--how they mediate the computations that subserve sensation, thought, emotion, and action, and how they are corrupted ...

Multiple-mouse Neuroanatomical Magnetic Resonance Imaging

The field of mouse phenotyping with magnetic resonance imaging (MRI) is rapidly growing, motivated by the need for improved tools for characterizing and evaluating mouse models of human disease. MRI is an excellent modality for investigating genetically altered animals. It is capable of whole brain coverage, can be used in vivo, and provides multiple contrast mechanisms for investigating different aspects of neuranatomy and physiology. The advent of high-field scanners along with the ability to scan multiple mice simultaneously allows for rapid phenotyping of novel mutations.
Effective mouse MRI studies require attention to many aspects of experiment design. In this article, we will describe general methods to acquire quality images for mouse phenotyping using a system that images mice concurrently in shielded transmit/receive radio frequency (RF) coils in a common magnet (Bock et al., 2003). We focus particularly on anatomical phenotyping, an important and accessible application that has shown a high potential for impact in many mouse models at our imaging centre. Before we can provide the detailed steps to acquire such images, there are important practical considerations for both in vivo brain imaging (Dazai et al., 2004) and ex vivo brain imaging (Spring et al., 2007) that should be noted. These are discussed below.

Magnetic resonance imaging (MRI) has become an increasingly popular tool for examining the phenotype of genetically altered mice. This article illustrates the methods necessary to achieve high-throughput phenotyping of genetically altered mice using multiple-mouse MRI.

The field of mouse phenotyping with magnetic resonance imaging (MRI) is rapidly growing, motivated by the need for improved tools for characterizing and ...

Imaging Pheromone Sensing in a Mouse Vomeronasal Acute Tissue Slice Preparation

Peter Karlson and Martin L&uuml;scher used the term pheromone for the first time in 19591 to describe chemicals used for intra-species communication. Pheromones are volatile or non-volatile short-lived molecules2 secreted and/or contained in biological fluids3,4, such as urine, a liquid known to be a main source of pheromones3. Pheromonal communication is implicated in a variety of key animal modalities such as kin interactions5,6, hierarchical organisations3 and sexual interactions7,8 and are consequently directly correlated with the survival of a given species9,10,11. In mice, the ability to detect pheromones is principally mediated by the vomeronasal organ (VNO)10,12, a paired structure located at the base of the nasal cavity, and enclosed in a cartilaginous capsule. Each VNO has a tubular shape with a lumen13,14 allowing the contact with the external chemical world. The sensory neuroepithelium is principally composed of vomeronasal bipolar sensory neurons (VSNs)15. Each VSN extends a single dendrite to the lumen ending in a large dendritic knob bearing up to 100 microvilli implicated in chemical detection16. Numerous subpopulations of VSNs are present. They are differentiated by the chemoreceptor they express and thus possibly by the ligand(s) they recognize17,18. Two main vomeronasal receptor families, V1Rs and V2Rs19,20,21,22, are composed respectively by 24023 and 12024 members and are expressed in separate layers of the neuroepithelium. Olfactory receptors (ORs)25 and formyl peptide receptors (FPRs)26,27 are also expressed in VSNs.
Whether or not these neuronal subpopulations use the same downstream signalling pathway for sensing pheromones is unknown. Despite a major role played by a calcium-permeable channel (TRPC2) present in the microvilli of mature neurons28 TRPC2 independent transduction channels have been suggested6,29. Due to the high number of neuronal subpopulations and the peculiar morphology of the organ, pharmacological and physiological investigations of the signalling elements present in the VNO are complex.
Here, we present an acute tissue slice preparation of the mouse VNO for performing calcium imaging investigations. This physiological approach allows observations, in the natural environment of a living tissue, of general or individual subpopulations of VSNs previously loaded with Fura-2AM, a calcium dye. This method is also convenient for studying any GFP-tagged pheromone receptor and is adaptable for the use of other fluorescent calcium probes. As an example, we use here a VG mouse line30, in which the translation of the pheromone V1rb2 receptor is linked to the expression of GFP by a polycistronic strategy.

In mice, the ability to detect pheromones is principally mediated by the vomeronasal organ (VNO). Here, an acute tissue slice preparation of VNO for performing calcium imaging is described. This physiological approach allows observations of subpopulations and/or individual neurons in a living tissue and is convenient for receptor-ligand identification.

Peter Karlson and Martin L&uuml;scher used the term pheromone for the first time in 19591 to describe chemicals used for intra-species communication. ...

A Procedure for Implanting a Spinal Chamber for Longitudinal In Vivo Imaging of the Mouse Spinal Cord

Studies in the mammalian neocortex have enabled unprecedented resolution of cortical structure, activity, and response to neurodegenerative insults by repeated, time-lapse in vivo imaging in live rodents. These studies were made possible by straightforward surgical procedures, which enabled optical access for a prolonged period of time without repeat surgical procedures. In contrast, analogous studies of the spinal cord have been previously limited to only a few imaging sessions, each of which required an invasive surgery. As previously described, we have developed a spinal chamber that enables continuous optical access for upwards of 8 weeks, preserves mechanical stability of the spinal column, is easily stabilized externally during imaging, and requires only a single surgery. Here, the design of the spinal chamber with its associated surgical implements is reviewed and the surgical procedure is demonstrated in detail. Briefly, this video will demonstrate the preparation of the surgical area and mouse for surgery, exposure of the spinal vertebra and appropriate tissue debridement, the delivery of the implant and vertebral clamping, the completion of the chamber, the removal of the delivery system, sealing of the skin, and finally, post-operative care. The procedure for chronic in vivo imaging using nonlinear microscopy will also be demonstrated. Finally, outcomes, limitations, typical variability, and a guide for troubleshooting are discussed.

A Procedure for Implanting a Spinal Chamber for Longitudinal In Vivo Imaging of the Mouse Spinal Cord

In this video, we describe a procedure for implanting a chronic optical imaging chamber over the dorsal spinal cord of a live mouse. The chamber, surgical procedure, and chronic imaging are reviewed and demonstrated.

Studies in the mammalian neocortex have enabled unprecedented resolution of cortical structure, activity, and response to neurodegenerative insults by ...

Whole-mount Imaging of Mouse Embryo Sensory Axon Projections

The visualization of full-length neuronal projections in embryos is essential to gain an understanding of how mammalian neuronal networks develop. Here we describe a method to label in situ a subset of dorsal root ganglion (DRG) axon projections to assess their phenotypic characteristics using several genetically manipulated mouse lines. The TrkA-positive neurons are nociceptor neurons, dedicated to the transmission of pain signals. We utilize a TrkAtaulacZ mouse line to label the trajectories of all TrkA-positive peripheral axons in the intact mouse embryo. We further breed the TrkAtaulacZ line onto a Bax null background, which essentially abolishes neuronal apoptosis, in order to assess growth-related questions independently of possible effects of genetic manipulations on neuronal survival. Subsequently, genetically modified mice of interest are bred with the TrkAtaulacZ/Bax null line and are then ready for study using the techniques described herein. This presentation includes detailed information on mouse breeding plans, genotyping at the time of dissection, tissue preparation, staining and clearing to allow for visualization of full-length axonal trajectories in whole-mount preparation.

We present here an optimized protocol to genotype, stain and prepare fetal mice for the imaging of peripheral nociceptor axon projections in the whole animal, as an effective method to assess sensory axon growth phenotypes in developing genetically engineered mice.

The visualization of full-length neuronal projections in embryos is essential to gain an understanding of how mammalian neuronal networks develop. Here we ...

3-D Imaging and Analysis of Neurons Infected In Vivo with Toxoplasma gondii

Toxoplasma gondii is an obligate, intracellular parasite with a broad host range, including humans and rodents. In both humans and rodents, Toxoplasma establishes a lifelong persistent infection in the brain. While this brain infection is asymptomatic in most immunocompetent people, in the developing fetus or immunocompromised individuals such as acquired immune deficiency syndrome (AIDS) patients, this predilection for and persistence in the brain can lead to devastating neurologic disease. Thus, it is clear that the brain-Toxoplasma interaction is critical to the symptomatic disease produced by Toxoplasma, yet we have little understanding of the cellular or molecular interaction between cells of the central nervous system (CNS) and the parasite. In the mouse model of CNS toxoplasmosis it has been known for over 30 years that neurons are the cells in which the parasite persists, but little information is available about which part of the neuron is generally infected (soma, dendrite, axon) and if this cellular relationship changes between strains. In part, this lack is secondary to the difficulty of imaging and visualizing whole infected neurons from an animal. Such images would typically require serial sectioning and stitching of tissue imaged by electron microscopy or confocal microscopy after immunostaining. By combining several techniques, the method described here enables the use of thick sections (160 &#181;m) to identify and image whole cells that contain cysts, allowing three-dimensional visualization and analysis of individual, chronically infected neurons without the need for immunostaining, electron microscopy, or serial sectioning and stitching. Using this technique, we can begin to understand the cellular relationship between the parasite and the infected neuron.

3-D Imaging and Analysis of Neurons Infected In Vivo with Toxoplasma gondii

Using this protocol, we were able to image 160 &#181;m thick brain sections from mice infected with the parasite Toxoplasma gondii, which enables visualization and analysis of the spatial relationship between the encysting parasite and the infected neuron.

Toxoplasma gondii is an obligate, intracellular parasite with a broad host range, including humans and rodents. In both humans and rodents, Toxoplasma ...

A Large Lateral Craniotomy Procedure for Mesoscale Wide-field Optical Imaging of Brain Activity

The craniotomy is a commonly performed procedure to expose the brain for in vivo experiments. In mouse research, most labs utilize a small craniotomy, typically 3 mm x 3 mm. This protocol introduces a method for creating a substantially larger 7 mm x 6 mm cranial window exposing most of a cerebral hemisphere over the mouse temporal and parietal cortices (e.g., bregma 2.5 - 4.5 mm, lateral 0 - 6 mm). To perform this surgery, the head must be tilted approximately 30&#176; and much of the temporal muscle must be retracted. Due to the large amount of bone removal, this procedure is intended only for acute experiments with the animal anesthetized throughout the surgery and experiment.
The main advantage of this innovative large lateral cranial window is to provide simultaneous access to both medial and lateral areas of the cortex. This large unilateral cranial window can be used to study the neural dynamics between cells, as well as between different cortical areas by combining multi-electrode electrophysiological recordings, imaging of neuronal activity (e.g., intrinsic or extrinsic imaging), and optogenetic stimulation. Additionally, this large craniotomy also exposes a large area of cortical blood vessels, allowing for direct manipulation of the lateral cortical vasculature.

This protocol presents a method for creating a large unilateral craniotomy over the temporal and parietal regions of the mouse cerebral cortex. This is especially useful for real time imaging over an expansive area of a cortical hemisphere.

The craniotomy is a commonly performed procedure to expose the brain for in vivo experiments. In mouse research, most labs utilize a small craniotomy, ...

Live Imaging of the Ependymal Cilia in the Lateral Ventricles of the Mouse Brain

Multiciliated ependymal cells line the ventricles in the adult brain. Abnormal function or structure of ependymal cilia is associated with various neurological deficits. The current ex vivo live imaging of motile ependymal cilia technique allows for a detailed study of ciliary dynamics following several steps. These steps include: mice euthanasia with carbon dioxide according to protocols of The University of Toledo&#8217;s Institutional Animal Care and Use Committee (IACUC); craniectomy followed by brain removal and sagittal brain dissection with a vibratome or sharp blade to obtain very thin sections through the brain lateral ventricles, where the ependymal cilia can be visualized. Incubation of the brain&#8217;s slices in a customized glass-bottom plate containing Dulbecco&#8217;s Modified Eagle&#8217;s Medium (DMEM)/High-Glucose at 37 &#176;C in the presence of 95%/5% O2/CO2 mixture is essential to keep the tissue alive during the experiment. A video of the cilia beating is then recorded using a high-resolution differential interference contrast microscope. The video is then analyzed frame by frame to calculate the ciliary beating frequency. This allows distinct classification of the ependymal cells into three categories or types based on their ciliary beating frequency and angle. Furthermore, this technique allows the use of high-speed fluorescence imaging analysis to characterize the unique intracellular calcium oscillation properties of ependymal cells as well as the effect of pharmacological agents on the calcium oscillations and the ciliary beating frequency. In addition, this technique is suitable for immunofluorescence imaging for ciliary structure and ciliary protein localization studies. This is particularly important in disease diagnosis and phenotype studies. The main limitation of the technique is attributed to the decrease in live motile cilia movement as the brain tissue starts to die.

Using high-resolution differential interference contrast (DIC) microscopy, an ex vivo observation of the beating of motile ependymal cilia located within the mouse brain ventricles is demonstrated by live-imaging. The technique allows a recording of the unique ciliary beating frequency and beating angle as well as their intracellular calcium oscillation pacing properties.

Multiciliated ependymal cells line the ventricles in the adult brain. Abnormal function or structure of ependymal cilia is associated with various ...

Time-lapse Confocal Imaging of Migrating Neurons in Organotypic Slice Culture of Embryonic Mouse Brain Using In Utero Electroporation

In utero electroporation is a rapid and powerful approach to study the process of radial migration in the cerebral cortex of developing mouse embryos. It has helped to describe the different steps of radial migration and characterize the molecular mechanisms controlling this process. To directly and dynamically analyze migrating neurons they have to be traced over time. This protocol describes a workflow that combines in utero electroporation with organotypic slice culture and time-lapse confocal imaging, which allows for a direct examination and dynamic analysis of radially migrating cortical neurons. Furthermore, detailed characterization of migrating neurons, such as migration speed, speed profiles, as well as radial orientation changes, is possible. The method can easily be adapted to perform functional analyses of genes of interest in radially migrating cortical neurons by loss and gain of function as well as rescue experiments. Time-lapse imaging of migrating neurons is a state-of-the-art technique that once established is a potent tool to study the development of the cerebral cortex in mouse models of neuronal migration disorders.

Time-lapse Confocal Imaging of Migrating Neurons in Organotypic Slice Culture of Embryonic Mouse Brain Using In Utero Electroporation

This protocol provides instructions for direct observation of radially migrating cortical neurons. In utero electroporation, organotypic slice culture, and time-lapse confocal imaging are combined to directly and dynamically study the effects of overexpression or downregulation of genes of interest in migrating neurons and to analyze their differentiation during development.

In utero electroporation is a rapid and powerful approach to study the process of radial migration in the cerebral cortex of developing mouse embryos. It ...

Recording Synaptic Plasticity in Acute Hippocampal Slices Maintained in a Small-volume Recycling-, Perfusion-, and Submersion-type Chamber System

Even though experiments on brain slices have been in use since 1951, problems remain that reduce the probability of achieving a stable and successful analysis of synaptic transmission modulation when performing field potential or intracellular recordings. This manuscript describes methodological aspects that might be helpful in improving experimental conditions for the maintenance of acute brain slices and for recording field excitatory postsynaptic potentials in a commercially available submersion chamber with an outflow-carbogenation unit. The outflow-carbogenation helps to stabilize the oxygen level in experiments that rely on the recycling of a small buffer reservoir to enhance the cost-efficiency of drug experiments. In addition, the manuscript presents representative experiments that examine the effects of different carbogenation modes and stimulation paradigms on the activity-dependent synaptic plasticity of synaptic transmission.

This protocol describes the stabilization of the oxygen level in a small volume of recycled buffer and methodological aspects of recording activity-dependent synaptic plasticity in submerged acute hippocampal slices.

Even though experiments on brain slices have been in use since 1951, problems remain that reduce the probability of achieving a stable and successful ...