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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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="https://www-jove-com.remotexs.ntu.edu.sg/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="https://www-jove-com.remotexs.ntu.edu.sg/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.

<table><tbody><tr><td>Vibratome Series 1000 Sectioning System</td><td>Technical Products International, Inc.</td><td></td><td>Other vibratomes are compatible</td></tr><tr><td>Glycerol</td><td>Fisher Scientific</td><td>BP229-1</td><td></td></tr><tr><td>Tween-20</td><td>Fisher Scientific</td><td>BP337-500</td><td></td></tr><tr><td>Premium Slides</td><td>Fisher Scientific</td><td>12-544-2</td><td></td></tr><tr><td>#1.5 Coverslips</td><td>VWR</td><td>48393 251</td><td></td></tr><tr><td>Diamond Scriber</td><td>VWR</td><td>52865-005</td><td></td></tr><tr><td>Zeiss LSM 510 Meta confocal microscope</td><td>Zeiss</td><td>LSM 510</td><td></td></tr><tr><td>Ketaject&amp;reg; Ketamine HCl Inj., USP 100 mg/ml</td><td>Western Medical Supply, Inc.</td><td>4165</td><td></td></tr><tr><td>AnaSed&amp;reg; Injection Xylazine 20 mg/ml</td><td>Lloyd Inc.</td><td></td><td></td></tr><tr><td>ZsGreen Mice</td><td>Jackson Laboratories</td><td>7906</td><td>B6.Cg-Gt(ROSA)26Sortm6(CAG-ZsGreen1)Hze/J</td></tr><tr><td>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><td>Human Foreskin Fibroblasts (HFF) cells</td><td></td><td></td><td>These are primary cells from human foreskins.&amp;nbsp; We make these in-house but they may be purchased from outside vendors.</td></tr><tr><td>Dulbecco&#039;s High Glucose Modified Eagles Medium (DMEM)</td><td>HyClone</td><td>SH30081.01</td><td></td></tr><tr><td>Penicillin Streptomycin Solution, 100x</td><td>Corning</td><td>30-002-Cl</td><td></td></tr><tr><td>200 mM L-alanyl-L-glutamine</td><td>Corning</td><td>25-015-Cl</td><td></td></tr><tr><td>25 cm&lt;sup&gt;2&lt;/sup&gt; Canted neck flask</td><td>Fisher Scientific</td><td>1012639</td><td></td></tr><tr><td>Phosphate-Buffered Saline, 1x Without Calcium and Magnesium</td><td>VWR</td><td>45000-446</td><td></td></tr><tr><td>Phosphate-Buffered Saline, 10x, USP Sterile Ultra Pure Grade</td><td>amresco</td><td>K813-500ml</td><td></td></tr><tr><td>Fetal Bovine Serum</td><td>Gibco</td><td>26140-079</td><td></td></tr><tr><td>Bright-Line Hemocytometer</td><td>Sigma-aldrich</td><td>Z359629-1EA</td><td></td></tr><tr><td>Mouse Brain Slicer Matrix</td><td>Zivic Instruments</td><td>BSMAS005-1</td><td></td></tr><tr><td>Sodium Chloride</td><td>Fisher Scientific</td><td>BP358-1</td><td></td></tr><tr><td>Heparin sodium salt from porcine intestinal mucosa</td><td>Sigma-aldrich</td><td>H3393-100KU</td><td></td></tr><tr><td>Paraformaldehyde</td><td>Fisher Scientific</td><td>O4042-500</td><td></td></tr><tr><td>20 ml Disposable Scintillation Vials</td><td>Fisher Scientific</td><td>FS74500-20</td><td></td></tr><tr><td>Alcohol, Ethyl, 95%, 190 Proof</td><td>In-house</td><td>17212945</td><td>This product is purchased from an in-house stockroom.&amp;nbsp; Other companies are compatible.</td></tr><tr><td>Imaris Software</td><td>Bitplane</td><td></td><td></td></tr><tr><td>Clear nail polish</td><td></td><td></td><td>Other brands are compatible</td></tr><tr><td>10 ml Syringe with Luer-Lok</td><td>VWR</td><td>BD309604</td><td>Other syringes are compatible</td></tr><tr><td>Three-way Stopcock</td><td></td><td></td><td>Any brand is compatible</td></tr><tr><td>Hypodermic needle</td><td></td><td></td><td>Any brand is compatible - used to pin down mouse.</td></tr><tr><td>Cell Scraper</td><td></td><td></td><td>Any brand is compatible</td></tr><tr><td>25 G x 12&quot; Tubing, Safety Blood Collection Set, with Luer Adapter</td><td>Greiner Bio-One</td><td>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.  Met dit protocol, konden we beeld 160 pm dikke hersencoupes van muizen geïnfecteerd met de parasiet Toxoplasma gondii, die visualisatie en analyse van de ruimtelijke relatie tussen de encysting parasiet besmette neuron maakt. 

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

A Thin-skull Window Technique for Chronic Two-photon In vivo Imaging of Murine Microglia in Models of Neuroinflammation

Traditionally in neuroscience, in vivo two photon imaging of the murine central nervous system has either involved the use of open-skull1,2 or thinned-skull 3 preparations. While the open-skull technique is very versatile, it is not optimal for studying microglia because it is invasive and can cause microglial activation. Even though the thinned-skull approach is minimally invasive, the repeated re-thinning of skull required for chronic imaging increases the risks of tissue injury and microglial activation and allows for a limited number of imaging sessions. Here we present a chronic thin-skull window method for monitoring murine microglia in vivo over an extended period of time using two-photon microscopy. We demonstrate how to prepare a stable, accessible, thinned-skull cortical window (TSCW) with an apposed glass coverslip that remains translucent over the course of three weeks of intermittent observation. This TSCW preparation is far more immunologically inert with respect to microglial activation than open craniotomy or repeated skull thinning and allows an arbitrary number of imaging sessions during a time period of weeks. We prepare TSCW in CX3CR1 GFP/+ mice 4 to visualize microglia with enhanced green fluorescent protein to &le;150 &mu;m beneath the pial surface. We also show that this preparation can be used in conjunction with stereotactic brain injections of the HIV-1 neurotoxic protein Tat, adjacent to the TSCW, which is capable of inducing durable microgliosis. Therefore, this method is extremely useful for examining changes in microglial morphology and motility over time in the living brain in models of HIV Associated Neurocognitive Disorder (HAND) and other neurodegenerative diseases with a neuroinflammatory component.

A Thin-skull Window Technique for Chronic Two-photon In vivo Imaging of Murine Microglia in Models of Neuroinflammation

We describe a method for repeatedly visualizing murine microglia and circulating monocytes in vivo over hours, days or weeks using transcranial two-photon microscopy. We demonstrate how to prepare a thinned-skull window that allows intermittent observation of quiescent microglia that can be activated by adjacent stereotactic injection of the HIV-1 regulatory protein Tat.

Traditionally in neuroscience, in vivo two photon imaging of the murine central nervous system has either involved the use of open-skull1,2 or ...

In Vivo Imaging Systems (IVIS) Detection of a Neuro-Invasive Encephalitic Virus

Modern advancements in imaging technology encourage further development and refinement in the way viral research is accomplished. Initially proposed by Russel and Burch in Hume's 3Rs (replacement, reduction, refinement), the utilization of animal models in scientific research is under constant pressure to identify new methodologies to reduce animal usage while improving scientific accuracy and speed. A major challenge to Hume's principals however, is how to ensure the studies are statistically accurate while reducing animal disease morbidity and overall numbers. Vaccine efficacy studies currently require a large number of animals in order to be considered statistically significant and often result in high morbidity and mortality endpoints for identification of immune protection. We utilized in vivo imaging systems (IVIS) in conjunction with a firefly bioluminescent enzyme to progressively track the invasion of the central nervous system (CNS) by an encephalitic virus in a murine model. Typically, the disease progresses relatively slowly, however virus replication is rapid, especially within the CNS, and can lead to an often, lethal outcome. Following intranasal infection of the mice with TC83-Luc, an attenuated Venezuelan equine encephalitis virus strain modified to expresses a luciferase gene; we are able to visualize virus replication within the brain at least three days before the development of clinical disease symptoms. Utilizing CNS invasion as a key encephalitic disease development endpoint we are able to quickly identify therapeutic and vaccine protection against TC83-Luc infection before clinical symptoms develop. With IVIS technology we are able to demonstrate the rapid and accurate testing of drug therapeutics and vaccines while reducing animal numbers and morbidity.

In Vivo Imaging Systems IVIS Detection of a Neuro-Invasive Encephalitic Virus

Utilizing luciferase and in vivo imaging systems (IVIS) as a novel means to identify disease endpoints before clinical developments occur. IVIS has allowed us to visualize in real time the invasion of encephalitic viruses over multiple days, providing a more accurate disease model for future study. It has also allowed us to identify the potential protective features of antivirals and vaccines faster than currently utilized animal models. The capability to utilize individual animals over multiple time points ensures reduced animal requirements, costs, and overall morbidity to the animals utilized ensuring a more humane and more scientific means of disease study.

Modern advancements in imaging technology encourage further development and refinement in the way viral research is accomplished. Initially proposed by ...

Immunology and Infection

Three-dimensional Imaging and Analysis of Mitochondria within Human Intraepidermal Nerve Fibers

The goal of this protocol is to study mitochondria within intraepidermal nerve fibers. Therefore, 3D imaging and analysis techniques were developed to isolate nerve-specific mitochondria and evaluate disease-induced alterations of mitochondria in the distal tip of sensory nerves. The protocol combines fluorescence immunohistochemistry, confocal microscopy and 3D image analysis techniques to visualize and quantify nerve-specific mitochondria. Detailed parameters are defined throughout the procedures in order to provide a concrete example of how to use these techniques to isolate nerve-specific mitochondria. Antibodies were used to label nerve and mitochondrial signals within tissue sections of skin punch biopsies, which was followed by indirect immunofluorescence to visualize nerves and mitochondria with a green and red fluorescent signal respectively. Z-series images were acquired with confocal microscopy and 3D analysis software was used to process and analyze the signals. It is not necessary to follow the exact parameters described within, but it is important to be consistent with the ones chosen throughout the staining, acquisition and analysis steps. The strength of this protocol is that it is applicable to a wide variety of circumstances where one fluorescent signal is used to isolate other signals that would otherwise be impossible to study alone.

This protocol uses three-dimensional (3D) imaging and analysis techniques to visualize and quantify nerve-specific mitochondria. The techniques are applicable to other situations where one fluorescent signal is used to isolate a subset of data from another fluorescent signal.

The goal of this protocol is to study mitochondria within intraepidermal nerve fibers. Therefore, 3D imaging and analysis techniques were developed to ...

Large-scale Three-dimensional Imaging of Cellular Organization in the Mouse Neocortex

The mammalian neocortex is composed of many types of excitatory and inhibitory neurons, each with specific electrophysiological and biochemical properties, synaptic connections, and in vivo functions, but their basic functional and anatomical organization from cellular to network scale is poorly understood. Here we describe a method for the three-dimensional imaging of fluorescently-labeled neurons across large areas of the brain for the investigation of the cortical cellular organization. Specific types of neurons are labeled by the injection of fluorescent retrograde neuronal tracers or expression of fluorescent proteins in transgenic mice. Block brain samples, e.g., a hemisphere, are prepared after fixation, made transparent with tissue clearing methods, and subjected to fluorescent immunolabeling of the specific cell types. Large areas are scanned using confocal or two-photon microscopes equipped with large working distance objectives and motorized stages. This method can resolve the periodic organization of the cell type-specific microcolumn functional modules in the mouse neocortex. The procedure can be useful for the study of three-dimensional cellular architecture in the diverse brain areas and other complex tissues.

Here we describe a procedure for tissue clearing, fluorescent labeling, and large-scale imaging of mouse brain tissue which, thereby, enables visualization of the three-dimensional organization of cell types in the neocortex.

The mammalian neocortex is composed of many types of excitatory and inhibitory neurons, each with specific electrophysiological and biochemical ...

Concomitant Isolation of Primary Astrocytes and Microglia for Protozoa Parasite Infection

Astrocytes and microglia are the most abundant glial cells. They are responsible for physiological support and homeostasis maintenance in the central nervous system (CNS). The increasing evidences of their involvement in the control of infectious diseases justify the emerging interest in the improvement of methodologies to isolate primary astrocytes and microglia in order to evaluate their responses to infections that affect the CNS. Considering the impact of Trypanosoma cruzi (T. cruzi) and Toxoplasma gondii (T. gondii) infection in the CNS, here we provide a method to extract, maintain, dissociate and infect murine astrocytes and microglia cells with protozoa parasites. Extracted cells from newborn cortices are maintained in vitro for 14 days with periodic differential media replacement. Astrocytes and microglia are obtained from the same extraction protocol by mechanical dissociation. After phenotyping by flow cytometry, cells are infected with protozoa parasites. The infection rate is determined by fluorescence microscopy at different time points, thus enabling the evaluation of differential ability of glial cells to control protozoan invasion and replication. These techniques represent simple, cheap and efficient methods to study the responses of astrocytes and microglia to infections, opening the field for further neuroimmunology analysis.

The overall goal of this protocol is to instruct how to extract, maintain, and dissociate murine astrocyte and microglia cells from the central nervous system, followed by infection with protozoa parasites.

Astrocytes and microglia are the most abundant glial cells. They are responsible for physiological support and homeostasis maintenance in the central ...

High-Resolution 3D Imaging of Rabies Virus Infection in Solvent-Cleared Brain Tissue

The visualization of infection processes in tissues and organs by immunolabeling is a key method in modern infection biology. The ability to observe and study the distribution, tropism, and abundance of pathogens inside of organ tissues provides pivotal data on disease development and progression. Using conventional microscopy methods, immunolabeling is mostly restricted to thin sections obtained from paraffin-embedded or frozen samples. However, the limited 2D image plane of these thin sections may lead to the loss of crucial information on the complex structure of an infected organ and the cellular context of the infection. Modern multicolor, immunostaining-compatible tissue clearing techniques now provide a relatively fast and inexpensive way to study high-volume 3D image stacks of virus-infected organ tissue. By exposing the tissue to organic solvents, it becomes optically transparent. This matches the sample&#8217;s refractive indices and eventually leads to a significant reduction of light scattering. Thus, in combination with long free working distance objectives, large tissue sections up to 1 mm in size can be imaged by conventional confocal laser scanning microscopy (CLSM) at high resolution. Here, we describe a protocol to apply deep-tissue imaging after tissue clearing to visualize rabies virus distribution in infected brains in order to study topics like virus pathogenesis, spread, tropism, and neuroinvasion.

Novel, immunostaining-compatible tissue clearing techniques like the ultimate 3D imaging of solvent-cleared organs allow the 3D visualization of rabies virus brain infection and its complex cellular environment. Thick, antibody-labeled brain tissue slices are made optically transparent to increase imaging depth and to enable 3D analysis by confocal laser scanning microscopy.

The visualization of infection processes in tissues and organs by immunolabeling is a key method in modern infection biology. The ability to observe and ...

Quantitative 3D Imaging of Trypanosoma cruzi-Infected Cells, Dormant Amastigotes, and T Cells in Intact Clarified Organs

Chagas disease is a neglected pathology that affects millions of people worldwide, mainly in Latin America. The Chagas disease agent, Trypanosoma cruzi (T. cruzi), is an obligate intracellular parasite with a diverse biology that infects several mammalian species, including humans, causing cardiac and digestive pathologies. Reliable detection of T. cruzi in vivo infections has long been needed to understand Chagas disease&#39;s complex biology and accurately evaluate the outcome of treatment regimens. The current protocol demonstrates an integrated pipeline for automated quantification of T. cruzi-infected cells in 3D-reconstructed, cleared organs. Light-sheet fluorescent microscopy allows for accurately visualizing and quantifying of actively proliferating and dormant T. cruzi parasites and immune effector cells in whole organs or tissues. Also, the CUBIC-HistoVision pipeline to obtain uniform labeling of cleared organs with antibodies and nuclear stains was successfully adopted. Tissue clearing coupled with 3D immunostaining provides an unbiased approach to comprehensively evaluate drug treatment protocols, improve the understanding of the cellular organization of T. cruzi-infected tissues, and is expected to advance discoveries related to anti-T. cruzi immune responses, tissue damage, and repair in Chagas disease.

Quantitative 3D Imaging of Trypanosoma cruzi-Infected Cells, Dormant Amastigotes, and T Cells in Intact Clarified Organs

The present protocol describes light-sheet fluorescent microscopy and automated software-assisted methods to visualize and precisely quantify proliferating and dormant Trypanosoma cruzi parasites and T cells in intact, cleared organs and tissues. These techniques provide a reliable way to evaluate treatment outcomes and offer new insights into parasite-host interactions.

Chagas disease is a neglected pathology that affects millions of people worldwide, mainly in Latin America. The Chagas disease agent, Trypanosoma cruzi ...

Live Animal Imaging and Cell Sorting Methods for Investigating Neurodegeneration in a C. elegans Excitotoxic Necrosis Model

Excitotoxic necrosis is a leading form of neurodegeneration. This process of regulated necrosis is triggered by the synaptic accumulation of the neurotransmitter glutamate, and the excessive stimulation of its postsynaptic receptors. However, information on the subsequent molecular events that culminate in the distinct neuronal swelling morphology of this type of neurodegeneration is lacking. Other aspects, such as changes in specific subcellular compartments, or the basis for the differential cellular vulnerability of distinct neuronal subtypes, remain under-explored. Furthermore, a range of factors that come into play in studies that use in vitro or ex vivo preparations might modify and distort the natural progression of this form of neurodegeneration. It is therefore important to study excitotoxic necrosis in live animals by monitoring the effects of interventions that regulate the extent of neuronal necrosis in the genetically amenable and transparent&#160;model system of the nematode Caenorhabditis elegans. This protocol describes methods of studying excitotoxic necrosis in C. elegans neurons, combining optical, genetic, and molecular analysis. To induce excitotoxic conditions in C. elegans, a&#160;knockout of a glutamate transporter gene (glt-3) is combined with a neuronal sensitizing genetic background (nuls5 [Pglr-1::G&#945;S(Q227L)]) to produce glutamate receptor hyperstimulation and neurodegeneration. Nomarski differential interference contrast (DIC), fluorescent, and confocal microscopy in live animals are methods used to quantify neurodegeneration, follow subcellular localization of fluorescently labeled proteins, and quantify mitochondrial morphology in the degenerating neurons. Neuronal Fluorescence Activated Cell Sorting (FACS) is used to distinctly sort at-risk neurons for cell-type specific transcriptomic analysis of neurodegeneration. A combination of live imaging and FACS methods as well as the benefits of the C. elegans model organism allow researchers to leverage this system to obtain reproducible data with a large sample size. Insights from these assays could translate to novel targets for therapeutic intervention in neurodegenerative diseases.

Live Animal Imaging and Cell Sorting Methods for Investigating Neurodegeneration in a C. elegans Excitotoxic Necrosis Model

In a C. elegans excitotoxicity model, this protocol employs in vivo imaging to analyze the regulation of necrotic neurodegeneration, the effect of genes encoding candidate mediators, and involvement of mitochondria. Cell dissociation and sorting is used to specifically obtain at-risk neurons for cell-specific transcriptomic analysis of neurodegeneration and neuroprotection mechanisms.

Excitotoxic necrosis is a leading form of neurodegeneration. This process of regulated necrosis is triggered by the synaptic accumulation of the ...

Utilizing In Vivo Postnatal Electroporation to Study Cerebellar Granule Neuron Morphology and Synapse Development

Neurons undergo dynamic changes in their structure and function during brain development to form appropriate connections with other cells. The rodent cerebellum is an ideal system to track the development and morphogenesis of a single cell type, the cerebellar granule neuron (CGN), across time. Here, in vivo electroporation of granule neuron progenitors in the developing mouse cerebellum was employed to sparsely label cells for subsequent morphological analyses. The efficacy of this technique is demonstrated in its ability to showcase key developmental stages of CGN maturation, with a specific focus on the formation of dendritic claws, which are specialized structures where these cells receive the majority of their synaptic inputs. In addition to providing snapshots of CGN synaptic structures&#160;throughout cerebellar development, this technique can be adapted to genetically manipulate granule neurons in a cell-autonomous manner to study the role of any gene of interest and its effect on CGN morphology, claw development, and synaptogenesis.

Utilizing In Vivo Postnatal Electroporation to Study Cerebellar Granule Neuron Morphology and Synapse Development

Here we describe a method to visualize synaptogenesis of granule neurons in the mouse cerebellum over the time course of postnatal brain development when these cells refine their synaptic structures and form synapses to integrate themselves into the overall brain circuit.

Neurons undergo dynamic changes in their structure and function during brain development to form appropriate connections with other cells. The rodent ...

Whole&#45;Organ Pathogen Tracking via Fluorescent Reporter Quantification

High&#45;Resolution Three&#45;Dimensional Whole&#45;Organ Tomography of Microbial Infections

Most infections take place within three-dimensional host tissues with intricate anatomy and locally varying host physiology. The positioning of pathogen cells within this diverse environment significantly affects their stress levels, responses, fate, and contribution to the overall progression of the disease and treatment failure. However, due to the technical difficulties in locating &#181;m-sized pathogen cells within cm-sized host organs, this area of research has been relatively unexplored. Here, we present a method for addressing this challenge. We employ serial two-photon tomography and AI-enhanced image analysis to locate individual Salmonella cells throughout the entire spleen, liver lobes, and whole lymph nodes of infected mice. Using fluorescent reporters and in vivo antibody administration, the replication rate of single Salmonella cells, their local interaction with specific immune cells, and bacterial responses to antibiotics can be determined. These methodologies open avenues for a comprehensive examination of infections, their prevention, and treatment within the three-dimensional tissue context.

Author Spotlight: Unraveling Bacterial Responses to Antibiotics and Immune System in Tissues

Here, we describe a procedure that enables organ-wide detection of pathogenic bacteria during infection and quantification of fluorescent reporter activities.

Most infections take place within three-dimensional host tissues with intricate anatomy and locally varying host physiology. The positioning of pathogen ...

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

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