The authors acknowledge Charles River laboratories for their excellent technical support executing the histopathology analyses. This work was funded by National Institute of Neurological Disorders and Stroke (NINDS) (RO1 NS033506 and RO1 NS060699). The funders had no role in study design, data collection and analysis, decision to publish, or preparation of the manuscript.

All animal experiments were performed in accordance to a protocol (number 2083-16 and 2083-19) reviewed and approved by the Institution Animal Care and Use Committee (IACUC) of Princeton University. This work was done by strictly following the biosafety level-2 (BSL-2) requirements, to which we have a fully equipped lab approved by the Princeton University biosafety committee. The procedures including mouse footpad abrasion, viral inoculation, mouse dissection, and tissue collection were performed in a biological safety cabinet (BSC) in the Princeton University biocontainment animal facility room. Those performing the procedure wore disposable gowns, head cover, eye protection, sterile gloves, surgical masks, and shoe covers.
1. Mouse footpad abrasion
<ol>
	<li>Anesthetize a C57BL/6 mouse (5&#8211;7 weeks old) with isoflurane gas anesthetic, delivered at a dosage of 3% via a small anesthesia system (chamber).
	<ol>
		<li>Place the mouse in the surgical plane of anesthesia on its back, prior to inoculation, in the hind footpad. Attach a nose cone with a diaphragm (a slit sufficiently sized to fit the muzzle of the animal) to the mouse and deliver isoflurane gas anesthetic at a constant dosage of 1.5%&#8211;2.0%.</li>
		<li>Closely monitor the mouse for a response to painful stimulus created by forceful pinching of the toe with forceps.</li>
	</ol>
	</li>
	<li>Lightly grasp one hind footpad with flat forceps.
	<ol>
		<li>Gently abrade the glabrous skin of the hind footpad approximately 20x, between the heel and walking pads, with an emery board (100&#8211;180 grit). Do not induce bleeding by abrading too frequently or applying too much pressure.</li>
		<li>Using fine forceps, slowly peel off the stratum corneum detached by abrasion to expose the stratum basale.</li>
	</ol>
	</li>
</ol>
2. PRV footpad inoculation
<ol>
	<li>Prepare the virus inoculum diluted to the desired titer in DMEM media containing 2% fetal bovine serum (FBS) and 1% penicillin/streptomycin (Table of Materials).
	<ol>
		<li>Quantitate the number of plaque-forming units (PFU) in the virus stock on PK15 cells to calculate and standardize the viral dose and dilute the viral inoculum accordingly. 
		NOTE: In this study, virulent strain of PRV (PRV-Becker) was used at a dose of 8 x 106 PFU. This dose was optimized in a previous preliminary experiment to ensure that all inoculated animals showed clinical symptoms at 82 h post-inoculation (hpi).</li>
		<li>Keep the virus inoculum on ice and gently mix before use.</li>
	</ol>
	</li>
	<li>Add a 20 &#956;L droplet of virus inoculum (8 x 106 PFU) onto the abraded footpad (topical administration). Carry out mock inoculations (medium only) in parallel.</li>
	<li>Gently rub 10x with the shaft of a needle to facilitate adsorption of the virus. Avoid scratching with the needle point. Repeat this step every 10 min.</li>
	<li>Keep the mouse under anesthesia for 30 min until the abraded footpad is dry.</li>
	<li>After stopping anesthesia, monitor the mouse until it can maintain sternal recumbency and place it in an individual cage for clinical follow-up and sampling.</li>
</ol>
3. Mouse dissection and tissue collection
<ol>
	<li>At appropriate times after infection, euthanize the mouse by the asphyxiation method (CO2). 
	NOTE: In this study, the humane endpoint (when animals begin to exhibit terminal symptoms) for PRV-Becker infected mice is 82 hpi. Euthanize control animals in parallel.</li>
	<li>Place the mouse ventral side facing up on a surgical mat using needles/pins. Secure the limbs of the mouse with pins to a surgical foam board. Wet the ventral side of the mouse with 70% ethanol to minimize fur contamination.</li>
	<li>Pinch the external layer of fur and skin using forceps and make a small initial incision using fine scissors near the urethral opening.
	<ol>
		<li>From this opening, continue the incision on the mid ventral side up to the chin.</li>
		<li>Extend two lateral incisions towards the extremities of the forelimbs and hind limbs.</li>
		<li>Separate the skin from the underlying muscular layer and pin down to the side.</li>
		<li>Open the abdominal cavity and incise up to the base of the thorax. Complete the opening of the abdominal cavity by making two transversal incisions.</li>
		<li>Open the thoracic cavity by cutting the diaphragm and ribs on both lateral sides. 
		NOTE: Cutting the sides of the ribcage prevents the risk of cutting the heart.</li>
		<li>Collect exposed organs, including heart, lungs, spleen, pancreas, liver, kidneys, and bladder in 1.5 mL microtubes. Keep the tubes on ice.</li>
		<li>Cut the abraded footpad between the heel and walking pads and place the piece of tissue in a tube as described in step 3.3.6.</li>
	</ol>
	</li>
	<li>Place the mouse dorsal side up and wet the fur with 70% ethanol. Cut down the skin layer to expose the vertebral column.
	<ol>
		<li>Make a small incision in the region of the pelvis. Strip the skin from the hind limbs towards the head.</li>
		<li>Remove the legs and arms by cutting with scissors parallel with and close to the spinal column on both sides.</li>
		<li>Cut the head off at the base of the skull (C1&#8211;C2 level).</li>
		<li>Cut open the skull with scissors from the foramen magnum to the frontal bone.</li>
		<li>Pull open the skull in lateral directions using forceps.</li>
		<li>Gently scoop out the brain using forceps and proceed as described in step 3.3.6.</li>
	</ol>
	</li>
	<li>Clean the spinal column by cutting muscle, fat and soft tissues using curved scissors. Cut the tail. Place the intact spinal column in a 15 mL tube containing sterile ice-cold phosphate-buffered saline (PBS). Keep tubes on ice until further dissection of the spinal cord and DRG.</li>
</ol>
4. Spinal cord and DRG extraction
NOTE: Extract spinal cord and DRG from the vertebrae column directly following mouse dissection. The protocol for spinal cord and DRG extraction has been adapted from a previous publication7.
<ol>
	<li>Remove the vertebrae column from ice-cold PBS tube and remove remaining soft tissues, which becomes easier after refrigeration.</li>
	<li>Make three transverse cuts in the column through the vertebrae (T1, L1, and S2 levels) using a razor blade. Place the three pieces in a 15 mm Petri dish containing sterile ice-cold PBS. Keep track and mark origin of segments for later analysis.</li>
	<li>Take one segment and secure it between forceps, dorsal side facing up. Make a single, longitudinal cut down through the midline using a razor blade, splitting the column in two equal halves.</li>
	<li>Place both halves in a new Petri dish containing new sterile ice-cold PBS. Ensure the correct orientation of the segment to be able to distinguish left and right sides.</li>
	<li>Under a dissecting microscope, gently peel the spinal cord out of the vertebrae column from each half in a rostral to caudal direction using fine forceps.</li>
	<li>Collect both spinal cord halves in 1.5 mL microtubes containing 500 &#956;L of sterile ice-cold PBS. Keep the tubes on ice.</li>
	<li>Gently remove the meninges from one side of the spinal column to the other to expose the DRG.</li>
	<li>Remove each DRG individually by grasping the exposed spinal nerve and pull it gently out of the spinal column. Place the collected DRG in a 15 mm Petri dish containing ice-cold PBS. Harvest all ipsilateral and contralateral DRG separately from the spinal column segment and proceed as described in step 4.5. 
	NOTE: The DRG is visible as a round transparent structure along the white spinal nerve.</li>
	<li>Repeat steps 4.3&#8211;4.7 using the two other spinal column segments within 30&#8211;45 min.</li>
	<li>Centrifuge all tubes at high speed (17,900 x g) for 3 min at 4 &#176;C.</li>
	<li>Aspirate supernatant and flash-freeze tubes in liquid nitrogen. Store tubes at -80 &#176;C for further tissue homogenization.</li>
	<li>Alternatively, fix and section the cleaned DRG and other mouse tissues for histopathological analyses and/or immunofluorescence staining after step 4.9.</li>
</ol>
5. Tissue homogenization
<ol>
	<li>Thaw the tubes containing frozen tissue samples on ice.</li>
	<li>Weigh 100 mg of tissue and place it in a 2 mL microcentrifuge tube containing a sterile steel bead and 500 &#956;L of RIPA buffer containing 0.5 M EDTA (pH = 8.0), 1 M Tris-HCl (pH = 8.0), 5 M NaCl, 10% SDS, and protease cocktail inhibitor tablets.</li>
	<li>Disrupt tissues at room temperature (RT) using a homogenizer at 20 cycles/s for 2 min, followed by a 1 min waiting period, then 20 cycles/s for 2 min.</li>
	<li>Centrifuge the tube at high speed (17,900 x g) for 10 min at RT.</li>
	<li>Collect supernatant (500 &#956;L) in a new tube and store it at -20 &#176;C until performing ELISA and qPCR to quantify inflammatory markers and viral loads, respectively. 
	NOTE: For qPCR, collect all samples with RNase-free material (i.e., beads, tubes, etc.) and disrupt tissues with RNA lysis buffer containing 1% betamercaptoethanol (Table of Materials).</li>
</ol>
6. Preparation and immunofluorescence staining of frozen DRG sections
<ol>
	<li>Tissue processing
	<ol>
		<li>Fix the dissected and cleaned DRG in 1% paraformaldehyde (PFA) for 2 h at RT.</li>
		<li>Transfer the tissue into a 15 mL tube with 10% sucrose in PBS. Incubate overnight at 4 &#176;C.</li>
		<li>Transfer the tissue into a 15 mL tube with 20% sucrose in PBS. Incubate overnight at 4 &#176;C.</li>
		<li>Transfer the tissue into a 15 mL tube with 30% sucrose in PBS. Incubate overnight at 4 &#176;C.</li>
		<li>Embed tissues in OCT using a cryomold and place blocks in dry ice for rapid freezing.</li>
		<li>Store the samples at -80 &#176;C until use.</li>
	</ol>
	</li>
	<li>Cryosectioning preparation
	<ol>
		<li>Remove the frozen block from -80 &#176;C and allow it to equilibrate in the cryostat chamber temperature for 30 min.</li>
		<li>Cryosection the DRG at a thickness of 15 &#181;m and mount the sections onto slides.</li>
		<li>Use a pen to draw a hydrophobic circle around the slide-mounted tissue.</li>
		<li>Keep the slides at 4 &#176;C until staining.</li>
	</ol>
	</li>
	<li>Immunofluorescence staining
	<ol>
		<li>Wash the DRG sections 3x in PBS for 10 min at RT.</li>
		<li>Incubate the sections with 100 &#181;L of desired primary antibody (e.g., mouse antibody against PRV glycoprotein B) for 1 h at 37 &#176;C. Dilute the primary antibody in PBS containing 10% negative goat serum.</li>
		<li>Repeat step 6.3.1.</li>
		<li>Incubate the sections with 100 &#181;L of desired secondary antibody (goat anti-mouse Alexa Fluor 488) for 50 min at 37 &#176;C. Dilute secondary antibody in PBS only.</li>
		<li>Add 100 &#181;L of DAPI (4&#8242;,6-diamidino-2-phenylindole) dye diluted in PBS onto the section. Further incubate samples 10 min at 37 &#176;C.</li>
		<li>Repeat step 6.3.1.</li>
		<li>Mount the samples using a fluorescence mounting medium onto glass slides and secure and cover with coverslip.</li>
	</ol>
	</li>
</ol>
7. Preparation and H&#38;E staining of paraffin-embedded tissue sections
<ol>
	<li>Fix the tissue in 10% formalin for 24 h at 4 &#176;C.</li>
	<li>Perform standard processing schedule for paraffin embedding and sectioning of fixed tissues. Transfer fixed tissues to 70% ethanol and process through upgraded alcohols to xylene followed by paraffin, as previously described8. 
	NOTE: Use polyester sponges to keep small tissue samples such as DRG inside cassettes.</li>
	<li>Perform standard deparaffinization followed by H&#38;E staining of tissue sections, as previously described9.</li>
</ol>

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The authors have nothing to disclose.

The footpad inoculation model described here is useful to investigate the initiation and development of neuroinflammatory responses during alphaherpesvirus infection. Moreover, this in vivo model is used to establish the kinetics of replication and spread of alphaherpesvirus from the PNS to CNS. This is an alternate to other inoculation models, such as the flank skin inoculation model, which relies on deep dermal scratching13, or the intracranial route, which directly introduces the virus into the CNS14,15,16. As a result, with the footpad model, it is possible to obtain a more detailed assessment of viral kinetics of replication and spread with associated local and distant immunopathological processes in the nervous system11,12.
In this protocol, the abrasion of the footpad and subsequent viral inoculation are crucial steps. Indeed, the stratum corneum needs to be completely removed after adequate abrasion in order to expose the stratum basale to viral inoculum for successful infection. However, the abrasion must be gentle and not induce bleeding, as this helps prevent infection of the blood circulation. The detached stratum corneum can be visualized by H&#38;E staining under light microscopy. The corneocytes present in the stratum corneum are flat, eosinophilic cells that lack nuclei. The volume of the viral inoculum (20 &#956;L droplet) has been optimized to ensure that the droplet stays on the footpad and covers the abraded site. Gently rubbing the inoculum droplet onto the abraded footpad is essential for efficient viral penetration. It is recommended to wait until the footpad is completely dry to stop the anesthesia and place the animal in a new cage. This step will avoid the mouse from licking the viral inoculum off the abraded footpad. It is recommended to process a maximum of three mice at once, using a nose cone that is set to expose them simultaneously to anesthesia.
While performing the mouse dissection, it is important to make cuts parallel to the vertebrae column, which prevents damage to the spinal cord and associated DRG. It also is suggested to remove as much fat, muscle, and soft tissue as possible to reduce accidental cutting into the spinal column and facilitate a better grip on the column segment with forceps before cutting down the midline.
It is also recommended to perform transverse cuts through the vertebrae column between discs in order to produce cleaned column segments and limit the risk of damaging DRG pairs. The meninges surrounding the spinal cord and covering the DRG must be completely removed to facilitate the identification and extraction of DRG. The DRG should be removed carefully from the spinal column without damage from the forceps. It is important that DRG remain intact for H&#38;E and immunofluorescence staining. Timing is critical for efficiency of this in vivo experiment, and the mouse dissection and spinal cord/DRG extraction should be performed consecutively in order to collect tissue that is as fresh as possible.
The tissue homogenization method described here has been optimized to ensure efficient disruption of a large number of heterogenous tissue samples. It is paramount to standardize the amount of tissue used, which allows direct comparison of ELISA and qPCR results between samples. For instance, it is suggested to weigh 100 mg of tissue for each homogenization procedure. Each tissue sample must be homogenized in its entirety, and leftovers should not be freeze-thawed. It is important to autoclave the steel beads and forceps before use to avoid any contamination during homogenization. A volume of 500 &#956;L is optimal to ensure complete homogenization of a 100 mg tissue sample. It is important to note that this limited volume allows only for the processing of three to four ELISA kits per sample.
Using the footpad inoculation model, it was demonstrated that PRV infection in mice induces severe inflammation, characterized by neutrophil infiltration in the footpad and DRG. High concentrations of inflammatory cytokines G-CSF and IL-6 were also detected in many homogenized tissues using ELISA. In addition, a strong correlation was found between PRV gene and protein expression (by qPCR and IF staining) in DRG and the production of both pro-inflammatory cytokines.
This model is suitable to compare the kinetics of viral replication/spread as well as neuroinflammatory responses among different alphaherpesvirus infections. For instance, regarding VZV, the restricted host-specificity and lack of clinical disease have limited the use of animal models17. Therefore, the mouse footpad PRV inoculation model may represent a new animal model for the study of the cellular and molecular mechanisms responsible for neuropathic pruritus in patients with post-herpetic lesions. Based on the similarities in clinical signs, pathogenesis, and genomes between VZV and PRV, it is believed that this mouse model will enhance the understanding of VZV pathogenesis and lead to the developments in innovative therapeutic strategies.
Finally, the model will guide research on peripheral neuropathies, such as multiple sclerosis and associated viral-induced damage to the PNS18. The pathogenesis of several neurotropic viruses (i.e., rabies virus, poliovirus, West Nile virus, Zika virus), which are known to infect the PNS, can also be studied using this model19,20,21,22. The footpad inoculation model can be used as a possible cost-effective tool for drug development. For instance, it may serve as a platform to screen and test the efficacy of anti-inflammatory and antiviral drugs designed to prevent viral-induced peripheral neuropathies.

This study describes a footpad inoculation model to investigate the replication and spread of viruses from the PNS to CNS and associated neuroinflammatory responses. The footpad inoculation model has been intensively used to study alphaherpesvirus infection in neurons1,2,3. The main objective of this model is to allow neurotropic viruses to travel a maximal distance through the PNS before reaching the CNS. Here, this model is used to obtain new insights in the development of a particular neuropathy (neuropathic itch) in mice infected with pseudorabies virus (PRV).
PRV is an alphaherpesvirus related to several well-known pathogens (i.e., herpes simplex type 1 and 2 [HSV1 and HSV2] and varicella-zoster virus [VZV]), which cause cold sores, genital lesions, and chicken pox, respectively4. These viruses are all pantropic and able to infect many different cell types without showing affinity for a specific tissue type. However, they all exhibit a characteristic neurotropism by invading the PNS (and occasionally, the CNS) of host species. The natural host is the pig, but PRV can infect most mammals. In these non-natural hosts, PRV infects the PNS and induces a severe pruritus called the &#8220;mad itch&#8221;, followed by peracute death5,6. The role of the neuroimmune response in the clinical outcome and pathogenesis of PRV infection has been poorly understood.
The footpad inoculation model allows PRV to initiate infection in the epidermal cells of the footpad. Then, the infection spreads into sensory and sympathetic nerve fibers that innervate the epidermis, sweat glands, and dermis. The infection spreads by virus particles moving via the sciatic nerve to the DRG within approximately 60 h. The infection spreads through the spinal cord, ultimately reaching the hindbrain when animals become moribund (82 h post-infection). During this time window, tissue samples can be collected, processed, and analyzed for virus replication and markers of the immune response. For instance, histological examination and viral load quantification can be performed in different tissues to establish correlations between the initiation and development of clinical, virological, and neuroinflammatory processes in PRV pathogenesis.
Using the footpad inoculation model, the cellular and molecular mechanisms of PRV-induced pruritus in mice can be investigated. Moreover, this model can provide new insight into the initiation and development of virus-induced neuroinflammation during herpesvirus infections. A better understanding of the processes underlying alphaherpesvirus-induced neuropathies may lead to the development of innovative therapeutic strategies. For instance, this model is useful to investigate the mechanisms of neuropathic itch in patients with post-herpetic lesions (e.g., herpes zoster, shingles) and test novel therapeutic targets in mice for the corresponding human diseases.

<table>
	<tbody>
		<tr>
			<td>Antibody anti-PRV gB</td>
			<td>Made by the lab</td>
			<td></td>
			<td>1/500 dilution</td>
		</tr>
		<tr>
			<td>Aqua-hold2 pap pen red</td>
			<td>Fisher scientific</td>
			<td>2886909</td>
			<td></td>
		</tr>
		<tr>
			<td>Compact emery boards-24 count (100/180 grit nail files)</td>
			<td>Revlon</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Complete EDTA-free Protease Inhibitor Cocktail</td>
			<td>Sigma-Aldrich</td>
			<td>11836170001</td>
			<td></td>
		</tr>
		<tr>
			<td>C57BL/6 mice (5-7 weeks)</td>
			<td>The Jackson Laboratories</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>DAPI solution (1mg/ml)</td>
			<td>Fisher scientific</td>
			<td>62248</td>
			<td>1/1000 dilution</td>
		</tr>
		<tr>
			<td>Disposable sterile polystyrene petri dish 100 x 15 mm</td>
			<td>Sigma-Aldrich</td>
			<td>P5731500</td>
			<td></td>
		</tr>
		<tr>
			<td>Dulbecco&#39;s Modified Eagle Medium (DMEM)</td>
			<td>Hyclone, GE Healthcare life Sciences</td>
			<td>SH30022</td>
			<td></td>
		</tr>
		<tr>
			<td>Dulbecco&#39;s Phophate Buffer Saline (PBS) solution</td>
			<td>Hyclone, GE Healthcare life Sciences</td>
			<td>SH30028</td>
			<td></td>
		</tr>
		<tr>
			<td>Fetal bovine serum (FBS)</td>
			<td>Hyclone, GE Healthcare life Sciences</td>
			<td>SH30088</td>
			<td></td>
		</tr>
		<tr>
			<td>Fine curved scissors stainless steel</td>
			<td>FST</td>
			<td>14095-11</td>
			<td></td>
		</tr>
		<tr>
			<td>Fluoromount-G mounting media</td>
			<td>Fisher scientific</td>
			<td>0100-01</td>
			<td></td>
		</tr>
		<tr>
			<td>Formalin solution, neutral buffered 10%</td>
			<td>Sigma-Aldrich</td>
			<td>HT501128</td>
			<td></td>
		</tr>
		<tr>
			<td>Isothesia Isoflurane</td>
			<td>Henry Schein</td>
			<td>NDC 11695-6776-2</td>
			<td></td>
		</tr>
		<tr>
			<td>Microcentrifuge tube 2ml</td>
			<td>Denville Scientific</td>
			<td>1000945</td>
			<td></td>
		</tr>
		<tr>
			<td>Microtube 1.5ml</td>
			<td>SARSTEDT</td>
			<td>72692005</td>
			<td></td>
		</tr>
		<tr>
			<td>Negative goat serum</td>
			<td>Vector</td>
			<td>S-1000</td>
			<td></td>
		</tr>
		<tr>
			<td>Penicillin/Streptomycin</td>
			<td>Gibco</td>
			<td>154022</td>
			<td></td>
		</tr>
		<tr>
			<td>Precision Glide needle 18G</td>
			<td>BD</td>
			<td>305196</td>
			<td></td>
		</tr>
		<tr>
			<td>Razor blades steel back</td>
			<td>Personna</td>
			<td>9412071</td>
			<td></td>
		</tr>
		<tr>
			<td>RNA lysis buffer (RLT)</td>
			<td>Qiagen</td>
			<td>79216</td>
			<td></td>
		</tr>
		<tr>
			<td>Stainless Steel Beads, 5 mm</td>
			<td>Qiagen</td>
			<td>69989</td>
			<td></td>
		</tr>
		<tr>
			<td>Superfrost/plus microscopic slides</td>
			<td>Fisher scientific</td>
			<td>12-550-15</td>
			<td></td>
		</tr>
		<tr>
			<td>Tissue lyser LT</td>
			<td>Qiagen</td>
			<td>69980</td>
			<td></td>
		</tr>
		<tr>
			<td>Tissue-Tek OCT</td>
			<td>Sakura</td>
			<td>4583</td>
			<td></td>
		</tr>
		<tr>
			<td>488 (goat anti-mouse)</td>
			<td>Life Technologies</td>
			<td>A11029</td>
			<td>1/2000 dilution</td>
		</tr>
	</tbody>
</table>

mouse footpad inoculation model to study viral-induced neuroinflammatory responses

This protocol describes a footpad inoculation model used to study the initiation and development of neuroinflammatory responses during alphaherpesvirus infection in mice. As alphaherpesviruses are main invaders of the peripheral nervous system (PNS), this model is suitable to characterize the kinetics of viral replication, its spread from the PNS to CNS, and associated neuroinflammatory responses. The footpad inoculation model allows virus particles to spread from a primary infection site in the footpad epidermis to sensory and sympathetic nerve fibers that innervate the epidermis, sweat glands, and dermis. The infection spreads via the sciatic nerve to the dorsal root ganglia (DRG) and ultimately through the spinal cord to the brain. Here, a mouse footpad is inoculated with pseudorabies virus (PRV), an alphaherpesvirus closely related to herpes simplex virus (HSV) and varicella-zoster virus (VZV). This model demonstrates that PRV infection induces severe inflammation, characterized by neutrophil infiltration in the footpad and DRG. High concentrations of inflammatory cytokines are subsequently detected in homogenized tissues by ELISA. In addition, a strong correlation is observed between PRV gene and protein expression (via qPCR and IF staining) in DRG and the production of pro-inflammatory cytokines. Therefore, the footpad inoculation model provides a better understanding of the processes underlying alphaherpesvirus-induced neuropathies and may lead to the development of innovative therapeutic strategies. In addition, the model can guide research on peripheral neuropathies, such as multiple sclerosis and associated viral-induced damage to the PNS. Ultimately, it can serve as a cost-effective in vivo tool for drug development.

The mouse footpad inoculation model allows for characterization of the immunopathogenesis of alphaherpesvirus infection in vivo, including replication and spread of the infection from the inoculated footpad to the nervous system and the induction of specific neuroinflammatory responses.
In this study, we first abraded the mouse hind footpad and either mock-inoculated or inoculated the abraded region with a virulent strain of PRV (PRV-Becker). The site of abrasion was visible in the control footpad. A crust was formed at the abrasion site as part of the healing process (Figure 1, black arrow). In contrast, mice inoculated with PRV showed severe inflammation at the humane endpoint (82 hpi), characterized by swelling of the footpad and redness.
Following footpad inoculation with the virulent PRV-Becker strain, mice began showing clinical signs at 72 hpi, characterized by swelling of the inoculated footpad and increasingly frequent tremors. By 82 hpi, PRV-Becker infected mice showed constant tremors in the inoculated leg and distinctive PRV symptoms, including intense scratching and biting of the foot. Severe inflammation was also observed in the footpad. The inflammatory response induced during PRV infection was then further characterized, including the infiltration of immune cells into tissues.
Histopathological examination of several tissues was performed, including the inoculated footpad and DRG, followed by H&#38;E staining of paraffin-embedded tissue sections. Epidermal necrosis and severe dermal inflammation (edema and fibrin) were observed in PRV-infected foot sections (Figure 2A, panel b). The epidermis, dermis, and connective tissues of infected mice showed a massive infiltration of neutrophils (identified by multilobed nuclei) marked with black arrows. Footpads of control mice were normal (Figure 2A, panel a). PRV-infected DRG showed minimal neuronal necrosis and mixed inflammation in infected mice while the DRG of control mice were normal (Figure 2B, panels a and b). The mixed inflammation infiltrate consisted mainly of neutrophils and lymphocytes.
Next, the kinetics of inflammatory cytokine production in mouse tissue after PRV footpad inoculation were established. Levels of specific inflammatory cytokines were quantified (i.e, interleukin-6 [IL-6] and granulocyte-colony stimulating factor [G-CSF]) from several tissues collected and homogenized from control and PRV-infected mice. The results demonstrated a significant increase of G-CSF levels in the footpad and DRG compared to controls at 7 hpi and 82 hpi (Figure 3A). Significant G-CSF levels were observed at 82 hpi in spinal cord, brain, heart, and liver tissue of PRV-infected mice compared to controls. Moreover, significant IL-6 levels were detected in all tissues of PRV-infected mice compared to controls starting at 24 hpi (Figure 3B).
The footpad inoculation model was further used to investigate PRV replication and spread from the inoculated footpad to the PNS and CNS and potential correlations with neuroinflammatory response development. PRV loads were quantified in several homogenized tissues by qPCR to amplify PRV DNA. DNA concentration was then converted to PFU, as previously described10. PRV loads were detected in the footpad starting at 24 hpi (~1 x 104 PFU/mg of tissue) and in DRG starting at 60 hpi (~1 x 103 PFU/mg of tissue; data not shown).
In a moribund state (82 hpi), PRV was detected in the footpad, DRG, spinal cord, and brain, with the highest concentration of PRV in DRG (~1 x 105 PFU/mg of tissue; Figure 4). PRV infection of DRG was confirmed by indirect immunofluorescence staining of DRG cryosections. PRV infection was detected in DRG neurons using anti-PRV gB antibody. PRV glycoprotein gB was expressed during late stages of infection in the cytoplasm of infected cells. As expected, the cytoplasmic expression of PRV gB (green) was confirmed in infected DRG, while no gB was expressed in control samples (Figure 5). Cell nuclei were identified with DAPI staining (blue).
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/61121/61121fig01.jpg" /> 
Figure 1: Representative images of mouse right hind paws after PRV inoculation.&#160;Mice are either mock-inoculated or inoculated with PRV in the abraded right hind footpad. PRV-inoculated footpad shows signs of inflammation, including redness and swelling at humane endpoint (82 hpi). The footpad of control mice appears normal with a dark red crust at the abraded site, indicating that the wound is healing. Black arrows indicate the site of abrasion. This figure has been modified from a previous publication11. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/61121/61121fig01large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/61121/61121fig02.jpg" /> 
Figure 2: Histopathological findings in footpad and DRG after PRV footpad inoculation.&#160;Hematoxylin and eosin (H&#38;E) staining of (A) mouse inoculated footpads and (B) and ipsilateral DRG from control (panel a) and PRV-infected (panel b) mice at 82 hpi. Histopathological manifestations observed in PRV-infected tissues (epidermal and neuronal necrosis and neutrophil infiltration) are absent from all examined mock-infected mice. Results are representative of three biological replicates for a given type of tissue. Black arrows indicate representative areas of inflammation with immune cell infiltration. Scale bars (50 &#956;m) are indicated for each picture. This figure has been modified from11. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/61121/61121fig02large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 3" class="xfigimg" src="/files/ftp_upload/61121/61121fig03.jpg" /> 
Figure 3: Kinetics of inflammatory cytokine production in homogenized mouse tissues after PRV footpad inoculation.&#160;(A) G-CSF and (B) IL-6 protein levels detected in PRV-infected (red) and control (black) mouse homogenized tissues at different hpi. Protein levels are quantified by ELISA and expressed as picogram (pg) per milligram (mg) of homogenized tissue (n = 5 per group, *p &#60; 0.05, ns = not significant). This figure has been modified from a previous publication12. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/61121/61121fig03large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 4" class="xfigimg" src="/files/ftp_upload/61121/61121fig04.jpg" /> 
Figure 4: Quantification of PRV genome in mouse homogenized tissues.&#160;PRV DNA is quantified in homogenized mouse tissues by qPCR using PRV UL54 primers. PRV loads are expressed as plaque forming units (PFU) per mg of tissue. PRV DNA is detected only in the foot, DRG, spinal cord, and brain (and not in other tissues), n = 10 per group. Dotted line shows the detection limit. This figure has been modified from a previous publication11. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/61121/61121fig04large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 5" class="xfigimg" src="/files/ftp_upload/61121/61121fig05.jpg" /> 
Figure 5: Assessment of PRV infection in DRG neurons by immunofluorescence staining.&#160;Confocal Z-stack images of mock- and PRV-infected DRG neurons after immunofluorescence staining using a mouse antibody specific for PRV gB (green). Cell nuclei are stained with DAPI (blue, panels a and c). Panel d shows several PRV-infected neurons expressing gB (white arrows). No gB expression is detected in the control DRG sections (panel b). Scale bars (50 &#956;m) are indicated for each picture. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/61121/61121fig05large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

Watch this Scientific Journal Video about Mouse Footpad Inoculation Model to Study Viral-Induced Neuroinflammatory Responses at JoVE.com

Mouse Footpad Inoculation Model to Study Viral-Induced Neuroinflammatory Responses

The footpad inoculation model is a valuable tool for characterizing viral-induced neuroinflammatory responses in vivo. In particular, it provides a clear assessment of viral kinetics and associated immunopathological processes initiated in the peripheral nervous system.

This protocol describes a footpad inoculation model used to study the initiation and development of neuroinflammatory responses during alphaherpesvirus ...

mouse-footpad-inoculation-model-to-study-viral-induced

Nanyang Technological University

Research

JoVE Journal

Immunology and Infection

10.5K Views.  Princeton University. The footpad inoculation model is a valuable tool for characterizing viral-induced neuroinflammatory responses in vivo. In particular, it provides a clear assessment of viral kinetics and associated immunopathological processes initiated in the peripheral nervous system.

Video: Mouse Footpad Inoculation Model to Study Viral-Induced Neuroinflammatory Responses

Protocols for Vaginal Inoculation and Sample Collection in the Experimental Mouse Model of Candida vaginitis

Vulvovaginal candidiasis (VVC), caused by Candida species, is a fungal infection of the lower female genital tract that affects approximately 75% of otherwise healthy women during their reproductive years18,32-34. Predisposing factors include antibiotic usage, uncontrolled diabetes and disturbance in reproductive hormone levels due to pregnancy, oral contraceptives or hormone replacement therapies33,34. Recurrent VVC (RVVC), defined as three or more episodes per year, affects a separate 5 to 8% of women with no predisposing factors33.
An experimental mouse model of VVC has been established and used to study the pathogenesis and mucosal host response to Candida3,4,11,16,17,19,21,25,37. This model has also been employed to test potential antifungal therapies in vivo13,24. The model requires that the animals be maintained in a state of pseudoestrus for optimal Candida colonization/infection6,14,23. Under such conditions, inoculated animals will have detectable vaginal fungal burden for weeks to months. Past studies show an extremely high parallel between the animal model and human infection relative to immunological and physiological properties3,16,21. Differences, however, include a lack of Candida as normal vaginal flora and a neutral vaginal pH in the mice.
Here, we demonstrate a series of key methods in the mouse vaginitis model that include vaginal inoculation, rapid collection of vaginal specimens, assessment of vaginal fungal burden, and tissue preparations for cellular extraction/isolation. This is followed by representative results for constituents of vaginal lavage fluid, fungal burden, and draining lymph node leukocyte yields. With the use of anesthetics, lavage samples can be collected at multiple time points on the same mice for longitudinal evaluation of infection/colonization. Furthermore, this model requires no immunosuppressive agents to initiate infection, allowing immunological studies under defined host conditions. Finally, the model and each technique introduced here could potentially give rise to use of the methodologies to examine other infectious diseases of the lower female genital tract (bacterial, parasitic, viral) and respective local or systemic host defenses.

Protocols for Vaginal Inoculation and Sample Collection in the Experimental Mouse Model of Candida vaginitis

Key techniques to be used in the evaluation of Candida vaginitis in an experimental animal model are described. The methods will allow rapid collection of vaginal specimens and lymphocytes from draining lumbar lymph nodes. These techniques could give rise to mouse models of other diseases in the female lower genital tract.

Vulvovaginal candidiasis (VVC), caused by Candida species, is a fungal infection of the lower female genital tract that affects approximately 75% of ...

Examining the Role of Nasopharyngeal-associated Lymphoreticular Tissue (NALT) in Mouse Responses to Vaccines

The nasopharyngeal-associated lymphoreticular tissues (NALT) found in humans, rodents, and other mammals, contribute to immunity in the nasal sinuses1-3. The NALT are two parallel bell-shaped structures located in the nasal passages above the hard palate, and are usually considered to be secondary components of the mucosal-associated lymphoid system4-6. Located within the NALT are discrete compartments of B and T lymphocytes interspersed with antigen-presenting dendritic cells4,7,8. These cells are surrounded by an epithelial cell layer intercalated with M-cells that are responsible for antigen retrieval from the mucosal surfaces of the air passages9,10. Naive lymphocytes circulating through the NALT are poised to respond to first encounters with respiratory pathogens7. While NALT disappear in humans by the age of two years, the Waldeyer's Ring and similarly structured lymphatic organs continue to persist throughout life6. In contrast to humans, mice retain NALT throughout life, thus providing a convenient animal model for the study of immune responses originating within the nasal sinuses11.
Cultures of single-cell suspensions of NALT are not practical due to low yields of mononuclear cells. However, NALT biology can be examined by ex vivo culturing of the intact organ, and this method has the additional advantage of maintaining the natural tissue structure. For in vivo studies, genetic knockout models presenting defects limited to NALT are not currently available due to a poor understanding of the developmental pathway. For example, while lymphotoxin-&alpha; knockout mice have atrophied NALT, the Peyer's patches, peripheral lymph nodes, follicular dendritic cells and other lymphoid tissues are also altered in these genetically manipulated mice12,13. As an alternative to gene knockout mice, surgical ablation permanently eliminates NALT from the nasal passage without affecting other tissues. The resulting mouse model has been used to establish relationships between NALT and immune responses to vaccines1,3. Serial collection of serum, saliva, nasal washes and vaginal secretions is necessary for establishing the basis of host responses to vaccination, while immune responses originating directly from NALT can be confirmed by tissue culture. The following procedures outline the surgeries, tissue culture and sample collection necessary to examine local and systemic humoral immune responses to intranasal (IN) vaccination.

Examining the Role of Nasopharyngeal-associated Lymphoreticular Tissue NALT in Mouse Responses to Vaccines

Methods to examine contributions of the nasopharyngeal-associated lymphoreticular tissues (NALT) to nasal and systemic immune responses of mice to intranasal vaccines are described. We demonstrate a surgical procedure to establish a NALT-dependent mouse model and ex vivo cultures of extracted NALT.

The nasopharyngeal-associated lymphoreticular tissues (NALT) found in humans, rodents, and other mammals, contribute to immunity in the nasal sinuses1-3. ...

An In vitro Model to Study Immune Responses of Human Peripheral Blood Mononuclear Cells to Human Respiratory Syncytial Virus Infection

Human respiratory syncytial virus (HRSV) infections present a broad spectrum of disease severity, ranging from mild infections to life-threatening bronchiolitis. An important part of the pathogenesis of severe disease is an enhanced immune response leading to immunopathology.	Here, we describe a protocol used to investigate the immune response of human immune cells to an HRSV infection. First, we describe methods used for culturing, purification and quantification of HRSV. Subsequently, we describe a human in vitro model in which peripheral blood mononuclear cells (PBMCs) are stimulated with live HRSV. This model system can be used to study multiple parameters that may contribute to disease severity, including the innate and adaptive immune response. These responses can be measured at the transcriptional and translational level. Moreover, viral infection of cells can easily be measured using flow cytometry. Taken together, stimulation of PBMC with live HRSV provides a fast and reproducible model system to examine mechanisms involved in HRSV-induced disease.

An In vitro Model to Study Immune Responses of Human Peripheral Blood Mononuclear Cells to Human Respiratory Syncytial Virus Infection

Human respiratory syncytial virus (HRSV) can cause severe bronchiolitis in young infants. Part of the pathogenesis of severe HRSV disease is caused by the host immune response. Stimulation of primary human immune cells with HRSV provides a fast and reproducible model system to study activation of inflammatory pathways and infection.

Human respiratory syncytial  virus (HRSV) infections present a broad spectrum of disease severity, ranging  from mild infections to life-threatening ...

Humanized Mouse Model to Study Bacterial Infections Targeting the Microvasculature

Neisseria meningitidis causes a severe, frequently fatal sepsis when it enters the human blood stream. Infection leads to extensive damage of the blood vessels resulting in vascular leak, the development of purpuric rashes and eventual tissue necrosis. Studying the pathogenesis of this infection was previously limited by the human specificity of the bacteria, which makes in vivo models difficult. In this protocol, we describe a humanized model for this infection in which human skin, containing dermal microvessels, is grafted onto immunocompromised mice. These vessels anastomose with the mouse circulation while maintaining their human characteristics. Once introduced into this model, N. meningitidis adhere exclusively to the human vessels, resulting in extensive vascular damage, inflammation and in some cases the development of purpuric rash. This protocol describes the grafting, infection and evaluation steps of this model in the context of N. meningitidis infection. The technique may be applied to numerous human specific pathogens that infect the blood stream.

Neisseria meningitidis is a human specific pathogen that infects blood vessels. In this protocol human microvessels are introduced into a mouse by grafting human skin onto immunocompromised mice. Bacteria adhere extensively to the human vessels, leading to vascular damage and development of the purpuric rash typically observed in human cases.

Neisseria meningitidis causes a severe, frequently fatal sepsis when it enters the human blood stream. Infection leads to extensive damage of the blood ...

Transplantation of Tail Skin to Study Allogeneic CD4 T Cell Responses in Mice

The study of T cell responses and their consequences during allo-antigen recognition requires a model that enables one to distinguish between donor and host T cells, to easily monitor the graft, and to adapt the system in order to answer different immunological questions. Medawar and colleagues established allogeneic tail-skin transplantation in mice in 1955. Since then, the skin transplantation model has been continuously modified and adapted to answer specific questions. The use of tail-skin renders this model easy to score for graft rejection, requires neither extensive preparation nor deep anesthesia, is applicable to animals of all genetic background, discourages ischemic necrosis, and permits chemical and biological intervention. 
In general, both CD4+ and CD8+ allogeneic T cells are responsible for the rejection of allografts since they recognize mismatched major histocompatibility antigens from different mouse strains. Several models have been described for activating allogeneic T cells in skin-transplanted mice. The identification of major histocompatibility complex (MHC) class I and II molecules in different mouse strains including C57BL/6 mice was an important step toward understanding and studying T cell-mediated alloresponses. In the tail-skin transplantation model described here, a three-point mutation (I-Abm12) in the antigen-presenting groove of the MHC-class II (I-Ab) molecule is sufficient to induce strong allogeneic CD4+ T cell activation in C57BL/6 mice. Skin grafts from I-Abm12 mice on C57BL/6 mice are rejected within 12-15 days, while syngeneic grafts are accepted for up to 100 days. The absence of T cells (CD3-/- and Rag2-/- mice) allows skin graft acceptance up to 100 days, which can be overcome by transferring 2 x 104 wild type or transgenic T cells. Adoptively transferred T cells proliferate and produce IFN-&#947; in I-Abm12-transplanted Rag2-/- mice.

Tail-skin transplantation is a powerful model for studying T cell-dependent rejection and tolerance induction during allogeneic immune responses in mice. The advantages of this protocol are minor invasive surgery, and ease of monitoring with no need to sacrifice the recipient mouse.

The study of T cell responses and their consequences during allo-antigen recognition requires a model that enables one to distinguish between donor and ...

Increased Recovery Time and Decreased LPS Administration to Study the Vagus Nerve Stimulation Mechanisms in Limited Inflammatory Responses

Inflammation is a local response to infection and tissue damage mediated by activated macrophages, monocytes, and other immune cells that release cytokines and other mediators of inflammation. For a long time, humoral and cellular mechanisms have been studied for their role in regulating the immune response, but recent advances in the field of immunology and neuroscience have also unraveled specific neural mechanisms with interesting therapeutic potential. The so-called cholinergic anti-inflammatory pathway (CAP) has been described to control innate immune responses and inflammation in a very potent manner. In the early 2000s, Tracey and collaborators developed a technique that stimulates the vagus nerve and mimics the effect of the pathway. The methodology is based on the electrical stimulation of the vagus nerve at low voltage and frequency, in order to avoid any side effects of overstimulation, such as deregulation of heart rate variability. Electrical devices for stimulation are now available, making it easy to set up the methodology in the laboratory. The goal of this research was to investigate the potential involvement of prostaglandins in the CAP. Unfortunately, based on earlier attempts, we failed to use the original protocol, as the induced inflammatory response either was too high or was not suitable for enzymatic metabolism properties. The different settings of the original surgery protocol remained mostly unchanged, but the conditions regarding inflammatory induction and the time point before sacrifice were improved to fit our purposes (i.e., to investigate the involvement of the CAP in more limited inflammatory responses). 
The modified version of the original protocol, presented here, includes a longer time range between vagus nerve stimulation and analysis, which is associated with a lower induction of inflammatory responses. Additionally, while decreasing the level of lipopolysaccharides (LPS) to inject, we also came across new observations regarding mechanistic properties in the spleen.

Vagus nerve stimulation has proven to have a strong efficacy for decreasing peripheral inflammation. Here, we present a modified vagus nerve stimulation protocol that allows for further examinations of the cholinergic anti-inflammatory mechanisms in limited inflammatory responses.

Inflammation is a local response to infection and tissue damage mediated by activated macrophages, monocytes, and other immune cells that release ...

An IL-8 Transiently Transgenized Mouse Model for the In Vivo Long-term Monitoring of Inflammatory Responses

Airway inflammation is often associated with bacterial infections and represents a major determinant of lung disease. The in vivo determination of the pro-inflammatory capabilities of various factors is challenging and requires terminal procedures, such as bronchoalveolar lavage and the removal of lungs for in situ analysis, precluding longitudinal visualization in the same mouse. Here, lung inflammation is induced through the intratracheal instillation of Pseudomonas aeruginosa culture supernatant (SN) in transiently transgenized mice expressing the luciferase reporter gene under the control of a heterologous IL-8 bovine promoter. Luciferase expression in the lung is monitored by in vivo bioluminescent image (BLI) analysis over a 2.5- to 48-h timeframe following the instillation. The procedure can be repeated multiple times within 2 - 3 months, thus permitting the evaluation of the inflammatory response in the same mice without the need to terminate the animals. This approach permits the monitoring of pro- and anti-inflammatory factors acting in the lung in real time and appears suitable for functional and pharmacological studies.

An IL-8 Transiently Transgenized Mouse Model for the In Vivo Long-term Monitoring of Inflammatory Responses

The method described here allows for the visualization of IL-8 promoter-dependent inflammation activation in the lungs of mice through non-invasive bioluminescence imaging (BLI). The same animal can be subjected to BLI multiple times for up to two months from the time of delivery of the luciferase reporter construct.

Airway inflammation is often associated with bacterial infections and represents a major determinant of lung disease. The in vivo determination of the ...

A Suction Blister Protocol to Study Human T-cell Recall Responses In Vivo

Cutaneous antigen-recall models allow for studies of human memory responses in vivo. When combined with skin suction blister (SB) induction, this model offers accessibility to rare populations of antigen-specific T-cells representative of the cellular memory response as well as the cytokine microenvironment in situ.
This report describes the practical procedure of a cutaneous recall, an SB induction, and a harvest of antigen-specific T-cells. To exemplify the method, the tuberculin skin test is used for antigenic recall in individuals who, prior to this study, underwent a Bacillus Calmette-Gu&#233;rin vaccination against an infection with Mycobacterium tuberculosis. Finally, examples of multiplex and flow cytometric analyses of SB specimens are provided, illustrating high fractions of antigen-specific polyfunctional CD4+ T-cells available by this sampling method compared with cells isolated from the blood.
The method described here is safe and minimally invasive, provides a unique opportunity to study both innate and adaptive immune responses in vivo, and may be beneficial to a broad community of researchers working with cell-mediated immunity and human memory responses, in the context of vaccine development.

A Suction Blister Protocol to Study Human T-cell Recall Responses In Vivo

Here, we provide a demonstration of the suction blister cutaneous recall model. The model allows a simple access to study human in vivo adaptive immune responses, for instance in the context of vaccine development.

Cutaneous antigen-recall models allow for studies of human memory responses in vivo. When combined with skin suction blister (SB) induction, this model ...

A Galleria mellonella Oral Administration Model to Study Commensal-Induced Innate Immune Responses

The investigation of the immunogenic potential of commensal bacteria on the host immune system is one essential component when studying intestinal host-microbe interactions. It is well established that different commensals exhibit a different potential to stimulate the host intestinal immune system. Such investigations involve vertebrate animals, especially rodents. Since increasing ethical concerns are linked with experiments involving vertebrates, there is a high demand for invertebrate replacements models. 
Here, we provide a Galleria mellonella oral administration model using commensal non-pathogenic bacteria and the possible assessment of the immunogenic potential of commensals on the G. mellonella immune system. We demonstrate that G. mellonella is a useful alternative invertebrate replacement model that allows the analysis of commensals with different immunogenic potential such as Bacteroides vulgatus and Escherichia coli. Interestingly, the bacteria exhibited no killing effect on the larvae, which is similar to mammals. The immune responses of G. mellonella were comparable with vertebrate innate immune responses and involve recognition of the bacteria and production of antimicrobial molecules. We propose that G. mellonella was able to restore previous microbiota balance, which is well known from healthy mammalian individuals. Although providing comparable innate immune responses in both G. mellonella and vertebrates, G. mellonella does not harbor an adaptive immune system. Since the investigated components of the innate immune system are evolutionary conserved, the model allows a prescreening and first analysis of bacterial immunogenic properties.

A Galleria mellonella Oral Administration Model to Study Commensal-Induced Innate Immune Responses

Here, we provide a detailed protocol for an oral administration model using Galleria mellonella larvae and how to characterize induced innate immune responses. Using this protocol, researchers without practical experience will be able to use the G. mellonella force-feeding method.

The investigation of the immunogenic potential of commensal bacteria on the host immune system is one essential component when studying intestinal ...

Tailoring In Vivo Cytotoxicity Assays to Study Immunodominance in Tumor-specific CD8+ T Cell Responses

Carboxyfluorescein succinimidyl ester (CFSE)-based in vivo cytotoxicity assays enable sensitive and accurate quantitation of CD8+ cytolytic T lymphocyte (CTL) responses elicited against tumor- and pathogen-derived peptides. They offer several advantages over traditional killing assays. First, they permit the monitoring of CTL-mediated cytotoxicity within architecturally intact secondary lymphoid organs, typically in the spleen. Second, they allow for mechanistic studies during the priming, effector and recall phases of CTL responses. Third, they provide useful platforms for vaccine/drug efficacy testing in a truly in vivo setting. Here, we provide an optimized protocol for the examination of concomitant CTL responses against more than one peptide epitope of a model tumor antigen (Ag), namely, simian virus 40 (SV40)-encoded large T Ag (T Ag). Like most other clinically relevant tumor proteins, T Ag harbors many potentially immunogenic peptides. However, only four such peptides induce detectable CTL responses in C57BL/6 mice. These responses are consistently arranged in a hierarchical order based on their magnitude, which forms the basis for TCD8 &#8220;immunodominance&#8221; in this powerful system. Accordingly, the bulk of the T Ag-specific TCD8 response is focused against a single immunodominant epitope while the other three epitopes are recognized and responded to only weakly. Immunodominance compromises the breadth of antitumor TCD8 responses and is, as such, considered by many as an impediment to successful vaccination against cancer. Therefore, it is important to understand the cellular and molecular factors and mechanisms that dictate or shape TCD8 immunodominance. The protocol we describe here is tailored to the investigation of this phenomenon in the T Ag immunization model, but can be readily modified and extended to similar studies in other tumor models. We provide examples of how the impact of experimental immunotherapeutic interventions can be measured using in vivo cytotoxicity assays.

Tailoring In Vivo Cytotoxicity Assays to Study Immunodominance in Tumor-specific CD8+ T Cell Responses

We describe here a flow cytometry-based in vivo killing assay that enables examination of immunodominance in cytotoxic T lymphocyte (CTL) responses to a model tumor antigen. We provide examples of how this elegant assay may be employed for mechanistic studies and for drug efficacy testing.

Carboxyfluorescein succinimidyl ester (CFSE)-based in vivo cytotoxicity assays enable sensitive and accurate quantitation of CD8+ cytolytic T lymphocyte ...

Mouse Footpad Inoculation Model to Study Viral-Induced Neuroinflammatory Responses

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Protocols for Vaginal Inoculation and Sample Collection in the Experimental Mouse Model of Candida vaginitis

Examining the Role of Nasopharyngeal-associated Lymphoreticular Tissue (NALT) in Mouse Responses to Vaccines

An In vitro Model to Study Immune Responses of Human Peripheral Blood Mononuclear Cells to Human Respiratory Syncytial Virus Infection

Humanized Mouse Model to Study Bacterial Infections Targeting the Microvasculature

Transplantation of Tail Skin to Study Allogeneic CD4 T Cell Responses in Mice

Increased Recovery Time and Decreased LPS Administration to Study the Vagus Nerve Stimulation Mechanisms in Limited Inflammatory Responses

An IL-8 Transiently Transgenized Mouse Model for the In Vivo Long-term Monitoring of Inflammatory Responses

A Suction Blister Protocol to Study Human T-cell Recall Responses In Vivo

A Galleria mellonella Oral Administration Model to Study Commensal-Induced Innate Immune Responses

Tailoring In Vivo Cytotoxicity Assays to Study Immunodominance in Tumor-specific CD8⁺ T Cell Responses