We thank the entire Koshy lab for helpful discussions. We thank Patty Jansma and the University of Arizona Neuroscience Department for advice and help with imaging. We also thank the Porreca lab for the use of their Vibratome. This research was supported by the US National Institutes of Health (NIH NS065116, A.A.K.).

NOTE: Mice were bred and maintained in a temperature and humidity controlled room with 12 hr reversed light/dark cycles with food and water available&#160;ad libitum at the University of Arizona. Experiments were conducted under guidelines and approval of the Institutional Animal Care and Use Committee of the University of Arizona. All efforts were made to minimize suffering. The Cre-reporter mice are on a C57BL/6 background11 and are commercially available.
1. Mouse Infection
NOTE: The method of mouse infection with Toxoplasma gondii described below has been used in studies previously published10&#8211;12.
<ol style="margin-left: 40px;">
	<li>Grow Toxoplasma strains in human foreskin fibroblasts (HFFs) cultured in Dulbecco&#39;s High Glucose Modified Eagles Medium (DMEM) supplemented with 10% FBS, 100 U/ml penicillin, 100 mg/ml streptomycin, and 2 mM L-glutamine (cDMEM) in a T-25 flask inside a 5% CO2 incubator until parasites have formed large parasitophorous vacuoles.</li>
	<li>Syringe-release parasites by scraping cells from the bottom of the flask and passing the resulting solution through a 25 gauge needle connected to a 3 ml syringe 2 times inside the flask then transfer all solution to a 5ml syringe case connected to a 27 gauge needle that has been placed upside down inside a 15 ml conical tube. Connect the plunger to the syringe and pass the parasite solution into the conical tube.</li>
	<li>Centrifuge the tube for 10 min at 300 x g. Aspirate the supernatant then resuspend the pellet in 4-6 ml sterile USP grade 1x Phosphate Buffered Saline (PBS) pH 7.4 (Stock Solution).</li>
	<li>Load a hemocytometer with 10 &#181;l of stock solution, wait 5-10 min for the parasites to settle then count the number of parasites in the four corner and center squares of the middle large square.</li>
	<li>Dilute parasites to appropriate inoculum size of 100K/200 &#181;l 1x PBS then inject mice intraperitoneally (i.p.) with a total volume of 200 &#181;l. 
	NOTE: For this procedure, mice were inoculated with 100K Type III (CEP)13 tachyzoites in 200 &#181;l 1x PBS. When infecting mice with other strains of Toxoplasma, concentration of inoculum may need to be adjusted to account for level of virulence.</li>
</ol>
2. Perfusion and Brain Harvesting
NOTE: This protocol was conducted at 21 days post infection (dpi) as this represents the time point at which peak numbers of cysts are found in the mouse brain, but other time points may be used. Size, number, location, and presence or absence of cysts depends on many factors including mouse strain, parasite strain type as well as inoculum size7,14&#8211;17. Investigators will need to individually determine the appropriate inoculum for the mouse and Toxoplasma strain he/she uses.
<ol style="margin-left: 40px;">
	<li>Setup all surgical harvesting tools (Figure 1).</li>
	<li>For each mouse, prepare and keep on ice: 20ml syringe filled with 0.9% Sodium Chloride (NaCl) + 10 U/ml Heparin in 1x sterile USP grade PBS (Heparin/Saline); 20 ml syringe filled with 4% Paraformaldehyde (PFA); Scintillation vial with 15 ml 4% PFA. For multiple mice, prepare a beaker filled with the entire amount of each solution needed for all mice then draw up a fresh syringe full of solution for each perfusion.</li>
	<li>Anesthetize the mouse i.p with 200 &#181;l/25 g of body weight of Ketamine/Xylazine cocktail (24 mg/ml &#38; 4.8 mg/ml respectively) on the day of interest. Monitor the mouse for level of anesthesia by checking toe pinch reflex. Continue with protocol only once the mouse shows no response to a firm toe pinch. 
	NOTE: Other methods of anesthesia may be used as long as the appropriate level of anesthesia is reached before proceeding with the protocol.</li>
	<li>Place the mouse in supine position on the covered foam platform. Pin all four limbs out at the feet then spray the chest and abdomen with 70% Ethanol (EtOH) (Figure 2A). Addition of several sterile paper towels under the mouse may be used to help absorb overflow of perfusion fluids.</li>
	<li>Lift skin just below the sternum with forceps then use scissors to make an incision. Pull skin back to expose the chest (Figure 2B).</li>
	<li>Lift xiphoid process and make an abdominal incision just below the diaphragm, exposing the liver which is bluntly dissected away from the diaphragm. Make two incisions, one on the left and one on the right through the diaphragm and the bottom of the ribs. Extend the left and right incisions approximately 1/2-2/3 way up the ribcage, which is then reflected back to open the chest cavity (Figure 2C). 
	NOTE: Take care not to cut any organs or major blood vessels during this procedure as doing so will compromise the quality of perfusion.</li>
	<li>Perfuse transcardially with 10-20 ml Heparin/Saline followed by 10-20 ml 4% PFA. 
	NOTE: If the perfusion is done properly, the liver will turn from red to tan (Figure 2D). If not done properly, blood vessels will be visible (Figure 2E).</li>
	<li>Turn the mouse over onto its abdomen, re-pin feet then spray head and neck with 70% EtOH. Lift skin from the base of the neck and make an incision. Remove skin to expose skull (Figure 2F). Decapitate head at the level of the shoulders. Remove the skull, gently remove the brain, and place it into a scintillation vial filled with 4% PFA (Figure 2G-H). 
	NOTE: If well perfused, the brain will appear white with no visible blood vessels (Figure 2I). If the brain is not well perfused, blood vessels will be visible (Figure 2J).</li>
	<li>Place the scintillation vial with brain in 4% PFA at 4 &#176;C overnight, covered from light. Rinse the brain in non-USP grade 1x PBS and transfer to a new scintillation vial filled with chilled 1x PBS. Store the brain in 1x PBS at 4 &#176;C, covered from light. 
	NOTE: Successful clearing and imaging has been accomplished with sections stored in 1x PBS for up to one month but it is best to section the brain within a few days as prolonged storage in PBS will reverse the PFA fixation and could lead to less than optimal images. Autofluorescence and increased blur have been noted in some sections imaged after storage in 1x PBS for one month or longer.</li>
</ol>
3. Brain Sectioning
<ol style="margin-left: 40px;">
	<li>Remove the brain from 1x PBS and place it into a brain matrix (Figure 3A) to allow for more precise and even cuts through the brain. Trim off the cerebellum and olfactory bulbs, if present, and then cut the brain coronally into 2 or 3 sections. 
	NOTE: Trimming and cutting of the brain may be done without the aid of a brain matrix. Number of sections cut coronally will depend on the cutting distance of the Vibratome being used. The brain can be sectioned in any orientation, though sagittal and coronal sections are most commonly used for neuroanatomical identification.</li>
	<li>Affix each brain section onto a specimen mounting block with cyanoacrylate. Affix a small block of 5% agarose gel behind the brain section for support (Figure 3B). 
	NOTE: Other support mechanisms may be used, but without support, the Vibratome may push on the brain causing uneven sectioning.</li>
	<li>Cut tissue into 160 &#181;m (working distance of the objective used for imaging) thick sections with the Vibratome set to a speed of 4 and amplitude of 9. 
	NOTE: The settings for speed and amplitude may be adjusted depending on the particular machine being used in order to obtain even, smooth sections. If the settings are not correct, the result can be uneven sections, tearing of tissue or chatter marks.</li>
	<li>Collect tissue sections into a scintillation vial filled with fresh, chilled 1x PBS if they are going to be cleared within one week. If sections are not going to be cleared within one week, place sections into 1.5 ml microcentrifuge tubes filled with cryopreservative solution and store at -20 &#176;C.</li>
</ol>
4. Optical Clearing of Brain Tissue Sections
NOTE: This protocol has been modified from a previously published method 8.
<ol style="margin-left: 40px;">
	<li>Prepare 25%, 50%, 75% and 90% vol/vol Glycerol in 1x PBS + 2% Tween-20 (Gl/PBST).</li>
	<li>Remove sections from 1x PBS or, if in cryopreservative, rinse in 1x PBS and transfer to a scintillation vial containing 10 ml of 25% Gl/PBST. Place vial on a shaker at 4 &#176;C, covered from exposure to light, for 12 hr.</li>
	<li>Confirm that all sections have sunk to the bottom of the vial. Aspirate the 25% Gl/PBST, leaving just enough to cover the sections then fill the vial with 10 ml 50% Gl/PBST and place on a shaker at 4 &#176;C, covered from exposure to light, for 12 hr. Repeat this process for the 75% Gl/PBST then the 90% Gl/PBST. Leave sections in the 90% Gl/PBST for 24 hr to reach maximal clearing. Over the course of the clearing process, observe gradual optical clearing (Figure 4). 
	NOTE: This process may be expedited if solutions are changed immediately after sections have sunk to the bottom of the vial. In 75% &#38; 90% solutions, sections will not sink all the way to the bottom of the vial but will float in the middle.</li>
</ol>
5. Mounting Brain Tissue Sections
<ol style="margin-left: 40px;">
	<li>Cut a No. 1.5 coverslip into 5 pieces to create a spacer. Use a ruler and diamond-tipped pencil to score and break the coverslip along the dashed lines shown in Figure 5A. Discard the center piece, arrange the 4 outer pieces on a plain glass slide to create a window, and then brush clear nail polish onto the seams to adhere the pieces of the spacer to the slide in the correct configuration (Figure 5B). 
	NOTE: The spacer will be used to create the necessary space between the slide and the coverslip for mounting the cleared sections. The coverslip may be cut in different ways to create any size window necessary. Pre-cut foam spacers are also commercially available. To allow nail polish to dry completely, it is best to make slides with spacers at least one day prior to mounting sections.</li>
	<li>Transfer sections to a slide using a plastic transfer pipet with the tip cut off. Make sure to fill in surrounding area with 90% Gl/PBST. Carefully lower a coverslip over the sections making sure not to introduce any bubbles. Excess Gl/PBST may be wicked away with a lint-free wipe.</li>
	<li>Image GFP+ neurons containing RFP+ Toxoplasma gondii cysts with a confocal microscope immediately (Figure 7A-B) then store slides flat and covered from light at 4 &#176;C. 
	NOTE: Alternatively, seal slides completely around all edges to store and imaged at a later time. Use of clear nail polish to seal slide is optional and may make it more difficult to remove coverslip if one were to remove sections from the slide at a later date for further processing.</li>
</ol>
6. Imaging
NOTE: Any microscope may be used that has the capability of obtaining high-resolution z-stacks.
<ol style="margin-left: 40px;">
	<li>After initially finding the focal plane at 10X, change to a 20X objective and scan through the section to identify a cyst located inside a GFP+ cell. Center the region of interest (ROI) in the field of view.</li>
	<li>Change to a 40X objective. Focus up and down through the entire ROI to make sure that the entire cell and cyst are visible. 
	NOTE: Images were obtained using a confocal microscope with a 40X/1.30 Oil DIC M27 objective.</li>
	<li>Using the installed imaging software, obtain a z-stack that includes the entire cell with cyst. Average settings used to obtain images are shown in Figure 6. 
	NOTE: Settings may need to be adjusted depending on each investigators tissue sections and microscope capabilities.</li>
	<li>Obtain a maximum projection image of the z-stack by setting the number of projections to 1. Obtain a full rotational view by setting the number of projections to 64. 
	NOTE: Other software packages are available and may be used to obtain maximum projection, rotational as well as other views of z-stack images.</li>
	<li>Analyze image with imaging analysis software.</li>
</ol>

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B. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Mice+infected+with+low-virulence+strains+of+Toxoplasma+gondii+lose+their+innate+aversion+to+cat+urine,+even+after+extensive+parasite+clearance.">Mice infected with low-virulence strains of Toxoplasma gondii lose their innate aversion to cat urine, even after extensive parasite clearance.</a> PloS one. 8 (9), 75246 (2013).</li><li>Evans, A. K., Strassmann, P. S., Lee, I. -. P., Sapolsky, R. 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The authors have nothing to disclose.

Given that cellular changes in infected host cells have been linked to disease outcomes in infections with other intracellular organisms such as HIV, Rabies, and Chlamydia18,19, we developed a technique that would allow us to study the intimate interactions that occur between the CNS host cell and Toxoplasma. The method described here accomplishes this goal by enabling efficient imaging of chronically infected neurons. Prior to the development of this method, such imaging was time consuming, costly, o...

A correction was made to <a href="http://www.jove.com/video/52237">"3-D Imaging and Analysis of Neurons Infected In Vivo with Toxoplasma gondii"</a>. The ordering of the authors was inverted. The order has been updated from:
 Anita A. Koshy, Carla M. Cabral
to:
Carla M. Cabral, Anita A. Koshy

A correction was made to <a href="http://www.jove.com/video/52237">"3-D Imaging and Analysis of Neurons Infected In Vivo with Toxoplasma gondii"</a>.

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

The overall goal of this method is to obtain high resolution, three-dimensional images of individual neurons that are infected by the obligate intracellular parasite Toxoplasma gondii.
Toxoplasma is often considered one of the most successful parasites because of its large intermediate host range, which includes humans and rodents. In both humans and rodents, after acute infection through ingestion of contaminated food or water, Toxoplasma is able to cause a persistent infection of the CNS by converting from its fast replicating form (the tachyzoite) to its slow-replicating and encysting form (the bradyzoite). In immunocompetent individuals, this latent CNS infection is thought to be relatively asymptomatic, but in immunocompromised individuals such as AIDS patients or transplant recipients, recrudescence of the parasite can lead to fatal toxoplasmic encephalitis1,2. In addition, recent studies have shown that latent infection with Toxoplasma can lead to behavioral changes in rodents3,4, though the mechanism remains unknown.
Surprisingly, despite these data highlighting the importance of the CNS-Toxoplasma interaction, relatively little is known about this relationship, especially at the cellular and molecular level. The ability to study even simple aspects of the brain-parasite interaction has been hampered in part by technologic limitations. For example, the majority of the work showing that neurons are the cells in which cysts persist has been done with electron microscopy (EM)5,6. Though EM gives high resolution, it is time consuming, labor intensive, and expensive. Immunofluorescence (IF) assays have recently been used in conjunction with confocal microscopy to confirm the work done by EM7. IF assays are technically easy to perform and relatively inexpensive, but using these techniques to understand the spatial relationship between the cyst and the infected neuron requires serial reconstruction, which is time consuming, technically difficult, and may lead to loss of valuable information. Thus, we have developed a method that can be used with the mouse model of CNS toxoplasmosis and allows us to image the entirety of infected neurons without EM or immunohistochemistry (IHC). By developing such a technique, we can begin to explore the cellular relationship between the infected cell and the cyst in a relatively quick and inexpensive manner.
The method we developed combines newer techniques for optically clearing and imaging thick brain sections by confocal microscopy8 with a system which marks in vivo cells that have been injected with parasite proteins9,10. In this system, we infect Cre-reporter mice that express a green fluorescent protein (GFP) only after Cre-mediated recombination11 with Toxoplasma strains that express a red fluorescent protein (RFP) and inject Cre recombinase into host cells9. This combination allows us to harvest the infected mouse brain after CNS infection is established, cut thick brain sections, and rapidly identify pertinent areas to image by finding the RFP+ cysts. It is important to note that as host cell expression of GFP depends solely on the injection of Cre by parasites, and not on infection, a number of the GFP+ cells do not contain parasites10. As the goal of this protocol is to be able to image whole infected neurons, the focus is only on GFP+ neurons that also contain an RFP+ cyst, but the protocol can also be used to image the GFP+/RFP- neurons.
Once the infected brain is harvested and sectioned, the sections are rendered transparent by glycerol clearing. Appropriate regions of sections are then imaged with confocal microscopy, allowing unprecedented visualization of infected host cells and the encysted parasites in their entirety. Here we provide a complete protocol for identifying, optically clearing, and imaging infected neurons.

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			<td height="21" style="height:21px;width:469px;">Name of Reagent/ Equipment</td>
			<td style="width:265px;">Company</td>
			<td style="width:135px;">Catalog Number</td>
			<td style="width:1008px;">Comments/Description</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Vibratome Series 1000 Sectioning System</td>
			<td>Technical Products International, Inc.</td>
			<td></td>
			<td>Other vibratomes are compatible</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Glycerol</td>
			<td>Fisher Scientific</td>
			<td>BP229-1</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Tween-20</td>
			<td>Fisher Scientific</td>
			<td>BP337-500</td>
			<td></td>
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			<td height="21" style="height:21px;">Premium Slides</td>
			<td>Fisher Scientific</td>
			<td>12-544-2</td>
			<td></td>
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			<td height="21" style="height:21px;">#1.5 Coverslips</td>
			<td>VWR</td>
			<td>48393 251</td>
			<td></td>
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		<tr height="21">
			<td height="21" style="height:21px;">Diamond Scriber</td>
			<td>VWR</td>
			<td>52865-005</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Zeiss LSM 510 Meta confocal microscope</td>
			<td>Zeiss</td>
			<td>LSM 510</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Ketaject&#174; Ketamine HCl Inj., USP 100mg/ml</td>
			<td>Western Medical Supply, Inc.</td>
			<td>4165</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">AnaSed&#174; Injection Xylazine 20mg/ml</td>
			<td>Lloyd Inc.</td>
			<td></td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">ZsGreen Mice</td>
			<td>Jackson Laboratories</td>
			<td>7906</td>
			<td>B6.Cg-Gt(ROSA)26Sortm6(CAG-ZsGreen1)Hze/J</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Surgical equipment</td>
			<td></td>
			<td></td>
			<td>Thumb forceps; Fine scissors-angled to side, sharp-sharp; Sharp-sharp scissors; Kelly hemostats; Mayo scissors; Micro spatula.</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Human Foreskin Fibroblasts (HFF) cells</td>
			<td></td>
			<td></td>
			<td>These are primary cells from human foreskins.&#160; We make these in-house but they may be purchased from outside vendors.</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Dulbecco&#39;s High Glucose Modified Eagles Medium (DMEM)</td>
			<td>HyClone</td>
			<td>SH30081.01</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:469px;">Penicillin Streptomycin Solution, 100X</td>
			<td>Corning</td>
			<td>30-002-Cl</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:469px;">200mM L-alanyl-L-glutamine</td>
			<td>Corning</td>
			<td>25-015-Cl</td>
			<td></td>
		</tr>
		<tr height="25">
			<td height="25" style="height:25px;width:469px;">25cm2 Canted neck flask</td>
			<td>Fisher Scientific</td>
			<td>1012639</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Phosphate-Buffered Saline, 1X Without Calcium and Magnesium</td>
			<td>VWR</td>
			<td>45000-446</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Phosphate-Buffered Saline, 10X, USP Sterile Ultra Pure Grade</td>
			<td>amresco</td>
			<td>K813-500ml</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:469px;">Fetal Bovine Serum</td>
			<td style="width:265px;">Gibco</td>
			<td style="width:135px;">26140-079</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:469px;">Bright-Line Hemocytometer</td>
			<td style="width:265px;">Sigma-aldrich</td>
			<td>Z359629-1EA</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Mouse Brain Slicer Matrix</td>
			<td>Zivic Instruments</td>
			<td>BSMAS005-1</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:469px;">Sodium Chloride</td>
			<td>Fisher Scientific</td>
			<td style="width:135px;">BP358-1</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:469px;">Heparin sodium salt from porcine intestinal mucosa</td>
			<td style="width:265px;">Sigma-aldrich</td>
			<td style="width:135px;">H3393-100KU</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:469px;">Paraformaldehyde</td>
			<td style="width:265px;">Fisher Scientific</td>
			<td style="width:135px;">O4042-500</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:469px;">20ml Disposable Scintillation Vials</td>
			<td style="width:265px;">Fisher Scientific</td>
			<td style="width:135px;">FS74500-20</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Alcohol, Ethyl, 95%, 190 Proof</td>
			<td style="width:265px;">In-house</td>
			<td>17212945</td>
			<td>This product is purchased from an in-house stockroom.&#160; Other companies are compatible.</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:469px;">Imaris Software</td>
			<td style="width:265px;">Bitplane</td>
			<td style="width:135px;"></td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:469px;">Clear nail polish</td>
			<td style="width:265px;"></td>
			<td style="width:135px;"></td>
			<td>Other brands are compatible</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:469px;">10ml Syringe with Luer-Lok</td>
			<td style="width:265px;">VWR</td>
			<td>BD309604</td>
			<td>Other syringes are compatible</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:469px;">Three-way Stopcock</td>
			<td style="width:265px;"></td>
			<td style="width:135px;"></td>
			<td>Any brand is compatible</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:469px;">Hypodermic needle</td>
			<td style="width:265px;"></td>
			<td style="width:135px;"></td>
			<td>Any brand is compatible - used to pin down mouse.</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:469px;">Cell Scraper</td>
			<td style="width:265px;"></td>
			<td style="width:135px;"></td>
			<td>Any brand is compatible</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">25G x 12&#34; Tubing, Safety Blood Collection Set, with Luer Adapter</td>
			<td style="width:265px;">Greiner Bio-One</td>
			<td style="width:135px;">450099</td>
			<td>Other brands are compatible</td>
		</tr>
	</tbody>
</table>

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.

Figure 7 includes representative images of two cyst-containing GFP+ neurons from two different 160 &#181;m thick sections as well as a representative measurement of the distance from cyst-to-cell-body for Figure 7B. Figures 7 A and B illustrate that this new protocol allows for visualization of the infected neuron in its entirety. Figure 7C shows that with this imaging technique, it is now possible to quantify the distance bet...

Watch this Scientific Journal Video about 3-D Imaging and Analysis of Neurons Infected In Vivo with Toxoplasma gondii at JoVE.com

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.

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 ...

3-d-imaging-analysis-neurons-infected-vivo-with-toxoplasma

Research

JoVE Journal

Neuroscience

11.4K Views.  University of Arizona.  Usando questo protocollo, siamo stati in grado di immagine di 160 micron di spessore sezioni di cervello di topi infettati con il parassita Toxoplasma gondii, che consente la visualizzazione e l'analisi del rapporto spaziale tra il parassita encysting e il neurone infetto. 

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

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 ...

Optical Imaging of Neurons in the Crab Stomatogastric Ganglion with Voltage-sensitive Dyes

Voltage-sensitive dye imaging of neurons is a key methodology for the understanding of how neuronal networks are organised and how the simultaneous activity of participating neurons leads to the emergence of the integral functionality of the network. Here we present the methodology of application of this technique to identified pattern generating neurons in the crab stomatogastric ganglion. We demonstrate the loading of these neurons with the fluorescent voltage-sensitive dye Di-8-ANEPPQ and we show how to image the activity of dye loaded neurons using the MiCAM02 high speed and high resolution CCD camera imaging system. We demonstrate the analysis of the recorded imaging data using the BVAna imaging software associated with the MiCAM02 imaging system. The simultaneous voltage-sensitive dye imaging of the detailed activity of multiple neurons in the crab stomatogastric ganglion applied together with traditional electrophysiology techniques (intracellular and extracellular recordings) opens radically new opportunities for the understanding of how central pattern generator neural networks work.

Here we present the methodology for fast and high resolution fluorescent voltage-sensitive dye imaging of detailed activity of neurons in the crab stomatogastric ganglion.

Voltage-sensitive dye imaging of neurons is a key methodology for the understanding of how neuronal networks are organised and how the simultaneous ...

Retrograde Fluorescent Labeling Allows for Targeted Extracellular Single-unit Recording from Identified Neurons In vivo

The overall goal of this method is to record single-unit responses from an identified population of neurons. In vivo electrophysiological recordings from individual neurons are critical for understanding how neural circuits function under natural conditions. Traditionally, these recordings have been performed 'blind', meaning the identity of the recorded cell is unknown at the start of the recording. Cellular identity can be subsequently determined via intracellular1, juxtacellular2 or loose-patch3 iontophoresis of dye, but these recordings cannot be pre-targeted to specific neurons in regions with functionally heterogeneous cell types. Fluorescent proteins can be expressed in a cell-type specific manner permitting visually-guided single-cell electrophysiology4-6. However, there are many model systems for which these genetic tools are not available. Even in genetically accessible model systems, the desired promoter may be unknown or genetically homogenous neurons may have varying projection patterns. Similarly, viral vectors have been used to label specific subgroups of projection neurons7, but use of this method is limited by toxicity and lack of trans-synaptic specificity. Thus, additional techniques that offer specific pre-visualization to record from identified single neurons in vivo are needed. Pre-visualization of the target neuron is particularly useful for challenging recording conditions, for which classical single-cell recordings are often prohibitively difficult8-11. The novel technique described in this paper uses retrograde transport of a fluorescent dye applied using tungsten needles to rapidly and selectively label a specific subset of cells within a particular brain region based on their unique axonal projections, thereby providing a visual cue to obtain targeted electrophysiological recordings from identified neurons in an intact circuit within a vertebrate CNS.
The most significant novel advancement of our method is the use of fluorescent labeling to target specific cell types in a non-genetically accessible model system. Weakly electric fish are an excellent model system for studying neural circuits in awake, behaving animals12. We utilized this technique to study sensory processing by "small cells" in the anterior exterolateral nucleus (ELa) of weakly electric mormyrid fish. "Small cells" are hypothesized to be time comparator neurons important for detecting submillisecond differences in the arrival times of presynaptic spikes13. However, anatomical features such as dense myelin, engulfing synapses, and small cell bodies have made it extremely difficult to record from these cells using traditional methods11, 14. Here we demonstrate that our novel method selectively labels these cells in 28% of preparations, allowing for reliable, robust recordings and characterization of responses to electrosensory stimulation.

Retrograde Fluorescent Labeling Allows for Targeted Extracellular Single-unit Recording from Identified Neurons In vivo

Retrograde transport of fluorescent dye labels a sub-population of neurons based on anatomical projection. Labeled axons can be visually targeted in vivo, permitting extracellular recording from identified axons. This technique facilitates recording when neurons cannot be labeled through genetic manipulation or are difficult to isolate using 'blind' in vivo approaches.

The overall goal of this method is to record single-unit responses from an identified population of neurons. In vivo electrophysiological recordings from ...

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

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

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

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

In Vivo Two-photon Imaging Of Experience-dependent Molecular Changes In Cortical Neurons

The brain's ability to change in response to experience is essential for healthy brain function, and abnormalities in this process contribute to a variety of brain disorders1,2. To better understand the mechanisms by which brain circuits react to an animal's experience requires the ability to monitor the experience-dependent molecular changes in a given set of neurons, over a prolonged period of time, in the live animal. While experience and associated neural activity is known to trigger gene expression changes in neurons1,2, most of the methods to detect such changes do not allow repeated observation of the same neurons over multiple days or do not have sufficient resolution to observe individual neurons3,4. Here, we describe a method that combines in vivo two-photon microscopy with a genetically encoded fluorescent reporter to track experience-dependent gene expression changes in individual cortical neurons over the course of day-to-day experience. 
One of the well-established experience-dependent genes is Activity-regulated cytoskeletal associated protein (Arc)5,6. The transcription of Arc is rapidly and highly induced by intensified neuronal activity3, and its protein product regulates the endocytosis of glutamate receptors and long-term synaptic plasticity7. The expression of Arc has been widely used as a molecular marker to map neuronal circuits involved in specific behaviors3. In most of those studies, Arc expression was detected by in situ hybridization or immunohistochemistry in fixed brain sections. Although those methods revealed that the expression of Arc was localized to a subset of excitatory neurons after behavioral experience, how the cellular patterns of Arc expression might change with multiple episodes of repeated or distinctive experiences over days was not investigated. 
In vivo two-photon microscopy offers a powerful way to examine experience-dependent cellular changes in the living brain8,9. To enable the examination of Arc expression in live neurons by two-photon microscopy, we previously generated a knock-in mouse line in which a GFP reporter is placed under the control of the endogenous Arc promoter10. This protocol describes the surgical preparations and imaging procedures for tracking experience-dependent Arc-GFP expression patterns in neuronal ensembles in the live animal. In this method, chronic cranial windows were first implanted in Arc-GFP mice over the cortical regions of interest. Those animals were then repeatedly imaged by two-photon microscopy after desired behavioral paradigms over the course of several days. This method may be generally applicable to animals carrying other fluorescent reporters of experience-dependent molecular changes4.

In Vivo Two-photon Imaging Of Experience-dependent Molecular Changes In Cortical Neurons

Experience-dependent molecular changes in neurons are essential for the brain's ability to adapt in response to behavioral challenges. An in vivo two-photon imaging method is described here that allows the tracking of such molecular changes in individual cortical neurons through genetically encoded reporters.

The brain's ability to change in response to experience is essential for healthy brain function, and abnormalities in this process contribute to a variety ...

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.

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 ...

In Vivo Multimodal Imaging and Analysis of Mouse Laser-Induced Choroidal Neovascularization Model

Laser-induced choroidal neovascularization (CNV) is a well-established model to mimic the wet form of age-related macular degeneration (AMD). In this protocol, we aim to guide the reader not simply through the technical considerations of generating laser-induced lesions to trigger neovascular processes, but rather focus on the powerful information that can be obtained from multimodal longitudinal in vivo imaging throughout the follow-up period.
The laser-induced mouse CNV model was generated by a diode laser administration. Multimodal in vivo imaging techniques were used to monitor CNV induction, progression and regression. First, spectral domain optical coherence tomography (SD-OCT) was performed immediately after the lasering to verify a break of Bruch&#39;s membrane. Subsequent in vivo imaging using fluorescein angiography (FA) confirmed successful damage of Bruch&#39;s membrane from serial images acquired at the choroidal level. Longitudinal follow-up of CNV proliferation and regression on days 5, 10, and 14 after the lasering was performed using both SD-OCT and FA. Simple and reliable grading of leaky CNV leasions from FA images is&#160;presented. Automated segmentation for measurement of total retinal thickness, combined with manual caliber application for measurement of retinal thickness at CNV sites, allow unbiased evaluation of the presence of edema. Finally, histological verification of CNV is performed using isolectin GS-IB4 staining on choroidal flatmounts. The staining is thresholded, and the isolectin-positive area is calculated with ImageJ.
This protocol is especially useful in therapeutics studies requiring high-throughput-like screening of CNV pathology as it allows fast, multimodal, and reliable classification of CNV pathology and retinal edema. In addition, high resolution SD-OCT enables the recording of other pathological hallmarks, such as the accumulation of subretinal or intraretinal fluid. However, this method does not provide a possibility to automate CNV volume analysis from SD-OCT images, which has to be performed manually.

In Vivo Multimodal Imaging and Analysis of Mouse Laser-Induced Choroidal Neovascularization Model

Here, we present the usefulness of longitudinal in vivo imaging in the follow-up of morphological changes of laser-induced choroidal neovascularization in mice.

Laser-induced choroidal neovascularization (CNV) is a well-established model to mimic the wet form of age-related macular degeneration (AMD). In this ...

In Vivo Two-photon Imaging of Cortical Neurons in Neonatal Mice

Two-photon imaging is a powerful tool for the in vivo analysis of neuronal circuits in the mammalian brain. However, a limited number of in vivo imaging methods exist for examining the brain tissue of live newborn mammals. Herein we summarize a protocol for imaging individual cortical neurons in living neonatal mice. This protocol includes the following two methodologies: (1) the Supernova system for sparse and bright labeling of cortical neurons in the developing brain, and (2) a surgical procedure for the fragile neonatal skull. This protocol allows the observation of temporal changes of individual cortical neurites during neonatal stages with a high signal-to-noise ratio. Labeled cell-specific gene silencing and knockout can also be achieved by combining the Supernova with RNA interference and CRISPR/Cas9 gene editing systems. This protocol can, thus, be used for analyzing the developmental dynamics of cortical neurons, molecular mechanisms that control the neuronal dynamics, and changes in neuronal dynamics in disease models.

In Vivo Two-photon Imaging of Cortical Neurons in Neonatal Mice

We present an in vivo two-photon imaging protocol for imaging the cerebral cortex of neonatal mice. This method is suitable for analyzing the developmental dynamics of cortical neurons, the molecular mechanisms that control the neuronal dynamics, and the changes in neuronal dynamics in disease models.

Two-photon imaging is a powerful tool for the in vivo analysis of neuronal circuits in the mammalian brain. However, a limited number of in vivo imaging ...

Generation of 3-D Collagen-based Hydrogels to Analyze Axonal Growth and Behavior During Nervous System Development

This protocol uses natural type I collagen to generate three-dimensional (3-D) hydrogel for monitoring and analyzing the axonal growth. The protocol is centered on culturing small pieces of embryonic or early postnatal rodent brains inside a 3-D hydrogel formed by the rat tail tendon-derived type I collagen with specific porosity. Tissue pieces are cultured inside the hydrogel and confronted to specific brain fragments or genetically-modified cell aggregates to produce and secrete molecules suitable for creating a gradient inside the porous matrix. The steps of this protocol are simple and reproducible but include critical steps to be considered carefully during its development. Moreover, the behavior of growing axons can be monitored and analyzed directly using a phase-contrast microscope or mono/multiphoton fluorescence microscope after fixation by immunocytochemical methods.

Here, we provide a method for analyzing the behavior of growing axons in 3D matrices, mimicking their natural development.

This protocol uses natural type I collagen to generate three-dimensional (3-D) hydrogel for monitoring and analyzing the axonal growth. The protocol is ...

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.