This work was funded by Candys Foundation.

All animal care and experimental procedures were performed in compliance with the European Community guide for the care and use of animals (2010/63/UE). Male C57BL/6JBomTac mice, aged 10 weeks, and male Sprague Dawley rats, aged 10 weeks, were used to demonstrate this protocol.
1. Preparation of the synthetic interstitial fluid
<ol>
	<li>Prepare synthetic interstitial fluid (SIF) according to the following recipe: 108 mM NaCl, 3.48 mM KCl, 3.50 mM MgSO4, 26 mM NaHCO3, 11.70 mM NaH2PO4, 1.50 mM CaCl2, 9.60 mM Na-gluconate, 5.50 mM glucose and 7.60 mM sucrose (see Table of Materials). 
	NOTE: SIF can be varied depending on the stimulation target, e.g., a calcium-free solution can be used when studying calcium channels.</li>
	<li>Adjust pH to 7.4 and stabilize the pH by carbogen gassing (5% CO2 and 95% O2)46.</li>
</ol>
2. Euthanasia
<ol>
	<li>Anesthetize adult mice and rats with a mixture of 70% CO2 and 30% O2. Decapitate mice using a pair of scissors and rats using a guillotine (see Table of Materials). 
	NOTE: Use a strain and age that fits the aim of the research. Both males and females&#160;can be used in this model.</li>
	<li>Separate the head from the body at the C3-C4 level of the spinal cord. 
	&#8203;NOTE: Euthanasia can also be performed with an intraperitoneal injection of pentobarbital (100-150 mg/kg).</li>
</ol>
3. Dissection
<ol>
	<li>Prepare the rat tissue following the steps below.
	<ol>
		<li>Remove the skin and the muscle around the head and neck using a pair of scissors (Figure 1A).</li>
		<li>Use a bone trimmer and scissors to separate the lower jaws from the head (Figure 1B-C).</li>
		<li>Open the spinal cord by inserting a bone trimmer caudally into the vertebrae&#39;s dorsal part and remove the vertebrae&#39;s dorsal part to expose the spinal cord and brainstem (Figure 1D).</li>
		<li>Cut the caudal part of the cranium by the borders of the occipital and interparietal bones to remove these bone structures exposing the cerebellum (Figure 1E). 
		NOTE: It is important not to damage the brainstem and spinal cord while cutting and removing the vertebras.</li>
		<li>Isolate the TNC (Sp5C) running caudally approximately 13-16 mm from the bregma on each side by cutting the dorsolateral part of the brainstem with spring scissors. Immerse the left and the right side TNC into SIF (Figure 1E-I). 
		NOTE: The description corresponds to adult rats.</li>
		<li>Cut the head mid-sagitally to divide the cranium in two using a saw (Figure 1J-L).</li>
		<li>Carefully remove the brain without touching the dura mater attached to the cranium using a spatula and cutting the trigeminal nerve where it enters the brainstem (Figure 1M-P).</li>
		<li>To isolate the TG, cut it, including its branches around the visual borders. Cut the mandibular branch where it enters the foramen ovale. Cut the ophthalmic and maxillary branches entering the skull as they are not divided macroscopically. While dissecting the TG, remove the dura mater covering the TG (Figure 1Q-T).</li>
		<li>Immerse the cranium halves and the TGs in SIF.</li>
	</ol>
	</li>
	<li>Prepare the mouse tissue following the steps below.
	<ol>
		<li>Remove skin and muscle around the head and neck using&#160;small scissors (Figure 2A-B).</li>
		<li>Open the spinal cord by inserting a pair of small scissors caudally into the vertebrae&#39;s dorsal part and remove the vertebrae&#39;s dorsal part to expose the spinal cord and brainstem (Figure 2C). 
		NOTE: It is important not to damage the spinal cord while cutting and removing the vertebras.</li>
		<li>Cut the cranium by the borders of the occipital and interparietal bones to remove these bone structures exposing the cerebellum (Figure 2D-F).</li>
		<li>Then, cut the parietal bone mid-sagitally and remove the bone to expose the cerebrum (Figure 2G-I).</li>
		<li>Carefully remove the cerebellum with a spatula to expose the brainstem (Figure 2J).</li>
		<li>Isolate the TNC-containing part of the brainstem using&#160;spring scissors (Figure 2K-N). Immerse the brainstem with TNC in SIF (Figure 2O).</li>
		<li>Remove the brain and cut the trigeminal nerve where it enters the brainstem (Figure 2P-Q).</li>
		<li>To isolate the TG, cut it, including its branches around the visual borders. Cut the mandibular branch where it enters the foramen ovale. Cut the ophthalmic and maxillary branches entering the skull as they are not divided macroscopically. While dissecting the TG, remove the dura mater covering the TG (Figure 2R-S).</li>
		<li>Immerse TGs in SIF (Figure 2T).</li>
	</ol>
	</li>
</ol>
4. Washing
<ol>
	<li>Wash skull halves, TNCs, and TGs in SIF for 30 min while replacing the SIF every 5 min at room temperature. 
	NOTE: The washing steps can be performed while keeping the tissue in plastic containers with a tulle lid to enable easy SIF exchange (Figure 3A-B).</li>
	<li>Prepare the rat tissue following the steps below.
	<ol>
		<li>Transfer TNC halves to separate microcentrifuge tube caps with 350 &#181;L of SIF (Figure 3C).</li>
		<li>Transfer the TGs to microcentrifuge tube caps - one TG per microcentrifuge tube cap with 350 &#181;L of SIF (Figure 3C).</li>
		<li>Place the skull halves on platforms made of clay or a 6-well culture plate and fill the skull with 400 &#181;L of SIF (Figure 3C).</li>
	</ol>
	</li>
	<li>Prepare the mouse tissue following the steps below.
	<ol>
		<li>Transfer the brainstem with TNC to a microcentrifuge tube cap with 250 &#181;L of SIF (Figure 3D).</li>
		<li>Transfer the two TGs to a microcentrifuge tube cap with 250 &#181;L of SIF (Figure 3D). 
		NOTE: Place two TGs per microcentrifuge tube cap when using tissue from mice.</li>
	</ol>
	</li>
	<li>Place rat skulls and microcentrifuge tube caps with rat and mouse tissue in a humidified incubator at 37 &#176;C. Replace SIF using a pipette every 5 min for 20 min. 
	&#8203;NOTE: It is important not to touch the tissue when adding and removing SIF.</li>
</ol>
5. Drug testing
<ol>
	<li>Determine the basal CGRP release levels.
	<ol>
		<li>Following the last wash, add 250 &#181;L of SIF to mouse TG and TNC. Add 350 &#181;L to rat TG and TNC and 400 &#181;L to each rat skull.</li>
		<li>After 10 min of incubation, collect 200 &#181;L of the sample in a microcentrifuge tube and add 50 &#181;L of 10x EIA buffer (provided with the CGRP enzyme immunoassay kit, see Table of Materials) to allow measurement of the basal CGRP release (step 6). Discard the remaining liquid. 
		NOTE: The incubation time must be the same for all samples.</li>
		<li>Immediately, store the samples at -20 &#176;C.</li>
		<li>After sampling of basal CGRP release levels, follow one of the three methods: (A) Single stimulation (step 5.2); (B) Concentration-response stimulation (step 5.3); and (C) Inhibition of stimulation (step 5.4) (Figure 4). 
		NOTE: The test compound and concentrations used depend on the aim of the study.</li>
	</ol>
	</li>
	<li>Perform single stimulation following the steps below.
	<ol>
		<li>Add test compound or vehicle to the tissue and leave it for 10 min (volume: 250 &#181;L for mouse TG and TNC, 350 &#181;L for rat TG and TNC, and 400 &#181;L for each rat skull).</li>
		<li>After 10 min of incubation, collect 200 &#181;L of the sample in a microcentrifuge tube with 50 &#181;L of 10x EIA buffer. Discard the remaining liquid and immediately store the samples at -20 &#176;C. 
		NOTE: The incubation time must be the same for all samples.</li>
	</ol>
	</li>
	<li>Perform concentration-response stimulation following the steps below.
	<ol>
		<li>Dilute the test compound or vehicle to the desired concentrations. Add the test compound in increasing concentrations starting with the lowest concentration. 
		NOTE: The test compound and concentrations used depend on the aim of the study. For example, 1 &#181;M, 10 &#181;M, and 100 &#181;M supercinnamaldehyde were used for the present study.</li>
		<li>Add the lowest concentration (1 &#181;M for the present study) of the test compound and corresponding vehicle to two identical tissue preparations and incubate for 10 min (volume: 250 &#181;L for mouse tissue, 350 &#181;L for rat TG and TNC, and 400 &#181;L for each rat skull).</li>
		<li>After 10 min of incubation, collect 200 &#181;L of the sample in a microcentrifuge tube with 50 &#181;L of 10x EIA buffer.</li>
		<li>Discard the remaining liquid and add the second lowest concentration (10 &#181;M for the present study) to the tissue.</li>
		<li>Immediately, store the samples at -20 &#176;C.</li>
		<li>Repeat this procedure with the remaining concentrations (100 &#181;M for the present study).</li>
	</ol>
	</li>
	<li>Perform inhibition of stimulation following the steps below.
	<ol>
		<li>Add the blocker or vehicle to the tissue and incubate for 10 min (volume: 250 &#181;L for mouse tissue, 350 &#181;L for rat TG and TNC, and 400 &#181;L for each rat skull). 
		NOTE: The blocker and concentration used depend on the aim of the study. For example, 3 &#181;M glibenclamide was used for the representative result in Figure 5.</li>
		<li>After 10 min of incubation, collect a 200 &#181;L sample in a microcentrifuge tube with 50 &#181;L of 10x EIA buffer. Discard the remaining liquid and immediately store the samples at -20 &#176;C.</li>
		<li>Add the agonist or agonist + blocker (see Table of Materials) to the tissue and incubate for 10 min. 
		NOTE: The agonist, blocker, and concentrations used depend on the aim of the study. In the present study, 3 &#181;M of glibenclamide and 1 &#181;M of capsaicin were used, Figure 5.</li>
		<li>After 10 min of incubation, collect a 200 &#181;L sample in a microcentrifuge tube with 50 &#181;L of 10x EIA buffer. Discard the remaining liquid and immediately store the samples at -20 &#176;C.</li>
	</ol>
	</li>
	<li>Perform a positive control for the experiment.
	<ol>
		<li>When appropriate, add a positive control (e.g., 1-10 &#181;M of capsaicin, see Table of Materials) to the tissue at the end of the protocol and collect a 200 &#181;L sample in a microcentrifuge tube with 50 &#181;L of 10x EIA buffer after an incubation period of 10 min. 
		NOTE: It is advantageous to include a positive control to ensure the setup and tissues are functioning. Capsaicin48,49,50 or the depolarizing stimulus of potassium (40-60 mM of KCl)46,47,49 are routinely used to cause the release of CGRP from the trigeminovascular system. 40-60 mM of KCl SIF is prepared as SIF, except NaCl is exchanged for KCl on an equimolar basis.</li>
	</ol>
	</li>
</ol>
6. Analysis of CGRP concentrations
<ol>
	<li>Measure the amount of CGRP released using an enzyme immunoassay (EIA) kit following the manufacturer&#39;s protocol (see Table of Materials).</li>
	<li>Measure the optical density at 410 nm using a plate photometer. If a different CGRP EIA kit is used, measure the optical density at the wavelength provided in the manufacturer&#39;s protocol. 
	NOTE: Samples must be diluted to match the standard curve.</li>
	<li>Perform data analysis.
	<ol>
		<li>Present the data either as absolute concentrations or normalize to the basal CGRP release from the specific tissue.</li>
	</ol>
	</li>
</ol>

The authors have nothing to disclose.

The method described was developed following studies showing the importance of CGRP in migraine pathophysiology. It is well suited for investigating the mechanisms involved in the release of CGRP from the trigeminovascular system, which is crucial for pain signaling in the head region. The amount of CGRP obtained in this model directly measures CGRP release from the trigeminal nerves innervating dura mater, TG, and TNC. The amount of CGRP release is quantitatively larger45,54 than the release measured in plasma after thermocoagulation in humans, trigeminal stimulation in cat39,55,56, and during migraine attacks23. One explanation could be that CGRP is diluted and degraded in the blood54. However, it is to be noted that direct stimulation with chemicals might be superior to pathophysiological activation. Further advantages are that it is possible to locate the release from three different sites within the trigeminovascular system and that it can be used together with pharmacological manipulation and in tissue from genetically modified rodents.
Lately, many preclinical rodent models focus on systemic administration of substances and subsequent pain or migraine-related read-outs using von Frey testing57,58, grimacing59,60,61 or light aversion62,63,64. These methods are useful in understanding the pain-inducing and pain-relieving properties of different substances. However, these approaches give no information on specific target tissues involved. In the present method, the trigeminovascular system is divided into three structures: the dura mater, TG, and TNC. This enables local exposure of each structure and the assessment of the location of action of a specific substance. This was utilized in a study from 2011, where the role of voltage-gated calcium channels in rats was explored, and the inhibition of these channels was found to be different in the three structures of the trigeminovascular pathway47. When dissecting structures of the nervous system, axotomy is unavoidable. Axotomy has been shown to alter the transcription of various genes65. These transcriptional changes are too slow to impact results from this method, but changes in phosphorylation cannot be excluded when compared to in vivo situations46. Although neuropeptides such as CGRP are formed in the cell soma, the release and action of neuropeptides are usually at the central or peripheral nerve terminals. Thus, studies of intact neurons, including terminals, are interesting when studying neuropeptide release. Therefore, methods of studying cultures of neurons from isolated ganglia have been established to serve as a model of the terminals. However, neuronal cell cultures are subject to several problems as mechanical dissociation can destroy neurons in a culture46. The longer time frame associated with culturing of cells leaves this method sensitive to transcriptomic changes due to axotomy and culture conditions65. Furthermore, the addition of growth factors and culturing on surface coatings have altered neuronal properties as transmitter and receptor expression66,67,68,69. These problems are avoided when studying freshly isolated intact ganglia instead of neuronal cell cultures.
One challenge with the ex vivo CGRP release method is the precise dissection of the tissue required for reproducible results. Especially accurate dissection of the TNC is challenging as this is a structure within the brainstem with no visible borders. Furthermore, the dura mater is fragile, and removal of the brain needs to be carefully carried out to ensure an intact structure. These obstacles can result in varying tissue size and, thus, varying basal and stimulation-induced CGRP levels. However, this variation can be accounted for by normalization to the basal CGRP release. It should also be noted that when isolating the TNC from mice, the entire lower part of the brainstem is isolated and not the more specific TNC-containing part as done in rats. In general, it can be an advantage to use rat tissue, as this allows measurement of CGRP release from dura mater and more precise dissection of the TNC. Furthermore, the size of the tissue also enables the use of a rat as its vehicle control, as one rat results in two cranium halves, two TGs, and two TNCs where one piece of the tissue is used for substance stimulation and the other for the vehicle. When using mice, two animals are needed for one experiment as both TGs are pooled in one sample, and the TNCs are dissected as one brainstem. Therefore, two TGs and one brainstem are used for substance stimulation, and two TGs and one brainstem from another mouse are used for vehicle control. This results in using twice as many mice compared to rats to obtain the same number of replicates. To reduce the number of mice used, a method to measure CGRP release from brainstem slices has been suggested49. It is an advantage that the method has been modified to enable the use of mice. This allows the use of many already available transgenic mouse strains, a useful tool for studying, e.g., signaling pathways. A positive control at the end of an experiment should be included to ensure that the tissue used in the experiment can release CGRP. The positive control could be TRPV1 agonist capsaicin or the depolarizing stimulus potassium (KCl), which have been found to release CGRP from the trigeminovascular system in both mice and rats46,47,48,49,50. Furthermore, the method has also been adapted to measure the release of other relevant peptides as pituitary adenylate cyclase-activating peptide (PACAP)&#160;- another peptide of great interest within migraine research70.
The method provides a useful tool for investigating CGRP release from specific target tissues in rats and mice. It is a relatively fast method that avoids problems associated with culturing neurons. The method protocol can easily be modified to study the concentration-response relationship or inhibition of a response by various pharmacological compounds. The ex vivo CGRP release method is one of the several preclinical methods useful for studying the role of CGRP and other mechanisms related to CGRP release in migraine pathophysiology.

Migraine is a neurological disorder that, according to WHO, is estimated to affect more than 1 billion people and is one of the leading causes of disability worldwide1. Thus, migraine has a significant impact on both patients and society. Despite the recent clinical success of CGRP antagonizing drugs, a large proportion of patients need improved treatment options2,3,4,5. Elucidation of migraine pathophysiology leading to novel effective treatments is required. Signaling within the trigeminovascular system consisting of the meninges, trigeminal ganglia (TG), and trigeminal nucleus caudalis (TNC) is central for migraine pathophysiology6,7.
The 37 amino acid neuropeptide calcitonin gene-related peptide (CGRP) was first discovered in the early 1980s when Amara and co-workers demonstrated that the primary RNA-transcript of the calcitonin gene could be processed to give mRNA coding for CGRP in addition to calcitonin8,9. The subsequent research suggested a link to migraine pathophysiology10. CGRP is a neurotransmitter with potent vasodilating properties11,12,13,14,15,16,17, and it is widely distributed in the central and peripheral nervous system13,14,18,19,20,21,22. The involvement of CGRP in migraine was underlined with the discovery of increased CGRP levels in the extracerebral circulation during migraine attacks in humans23, and that infusion of CGRP causes migraine-like pain in patients24. Two years later, the first proof-of-concept study of the effectivity of the CGRP antagonist olcegepant in treating migraine was published25.
CGRP is abundant in the trigeminovascular system as demonstrated in the TG21,26, sensory nerve fibers innervating the dura mater27,28,29, and TNC30. In the trigeminovascular system, CGRP is found in the small-to-medium-sized neurons of the TG, in unmyelinated C-fibers, and is expressed in nearly 50% of the neuronal population of the TG. The CGRP receptor is expressed mainly in larger neurons and found in myelinated A&#948;-fibers31,32. CGRP is released from neurons upon chemical or electrical stimulation33,34. Studies of pathways leading to the release of CGRP and the location of this activation are crucial to understand migraine pathophysiology. Throughout the last 5 decades, preclinical studies have contributed to gaining extensive knowledge on migraine-related signaling and have contributed to development of new treatments35. Many methods considering the vascular and neurogenic involvement have been modified and applied in migraine research. In vivo and in vitro models of arterial responses to biological compounds or pharmacological treatments17,36,37, and electrical nerve-stimulation38,39 can be mentioned. Furthermore, activated neurons in the TNC can be detected by c-Fos expression40,41,42 and electrophysiological recordings in this area43,44. Both methods measure nociceptive signals transmitted to the brain from the head, e.g., dura mater. The use of only one preclinical model does not present the full picture of migraine pathophysiology. Therefore, it is important to combine different models covering as many aspects of migraine pathophysiology as possible. The continued development of new models will cover various aspects of migraine mechanisms, and in time the mystery of migraine pathophysiology will be uncovered.
Here, a detailed protocol is presented of the CGRP release method, performed ex vivo in isolated TG and TNC from mice after chemical stimulation. CGRP release can also be studied in the dura mater from rats. Thus, in the experimental protocol for rats, dura mater is described together with TG and TNC. The basis for the CGRP release method was first described in 1999, where Ebersberger and co-workers conducted pioneering research and found that CGRP was released from dura mater after chemical and electrical stimulation of dural afferents in rats45. Later, this approach was extended to CGRP release from the TG46 and the TNC47. Subsequently, the method was modified to apply to TG and TNC in mice. So far, CGRP release from the dura mater has been challenging in mice.

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>6-well culture plate</td>
			<td>NUNC</td>
			<td>140675</td>
			<td></td>
		</tr>
		<tr>
			<td>Calcium chloride dihydrate</td>
			<td>Merck</td>
			<td>1.02382.1000</td>
			<td>For SIF buffer</td>
		</tr>
		<tr>
			<td>Caps for plastic containers</td>
			<td>ThermoFisher Scientific</td>
			<td>536617</td>
			<td></td>
		</tr>
		<tr>
			<td>Capsaicin</td>
			<td>Merck</td>
			<td>M2028</td>
			<td></td>
		</tr>
		<tr>
			<td>CGRP kits</td>
			<td>AH Diagnostics</td>
			<td>A05482.96</td>
			<td></td>
		</tr>
		<tr>
			<td>CO2</td>
			<td>Strandm&#248;llen</td>
			<td>4.6</td>
			<td>For carbogen gassing of SIF</td>
		</tr>
		<tr>
			<td>Delicate Bone Trimmer</td>
			<td>Fine Science Tools</td>
			<td>16109-14</td>
			<td></td>
		</tr>
		<tr>
			<td>Glibenclamide</td>
			<td>Tocris</td>
			<td>911</td>
			<td></td>
		</tr>
		<tr>
			<td>Glucose</td>
			<td>Merck</td>
			<td>G7021</td>
			<td>For SIF buffer</td>
		</tr>
		<tr>
			<td>Guillotine for rats</td>
			<td>Scandidact</td>
			<td>NS-802</td>
			<td></td>
		</tr>
		<tr>
			<td>Magnesium sulfate heptahydrate</td>
			<td>Merck</td>
			<td>M5921</td>
			<td>For SIF buffer</td>
		</tr>
		<tr>
			<td>Microcenrifuge tubes + lids/caps</td>
			<td>VWR</td>
			<td>700-5239</td>
			<td></td>
		</tr>
		<tr>
			<td>Mini Hacksaw</td>
			<td>BAHCO</td>
			<td>208</td>
			<td></td>
		</tr>
		<tr>
			<td>O2</td>
			<td>Strandm&#248;llen</td>
			<td>4.5</td>
			<td>For carbogen gassing of SIF</td>
		</tr>
		<tr>
			<td>Pentobarbital</td>
			<td>Glostrup pharmacy</td>
			<td>NA</td>
			<td>Magistral formula</td>
		</tr>
		<tr>
			<td>Plastic containers</td>
			<td>ThermoFisher Scientific</td>
			<td>536455</td>
			<td></td>
		</tr>
		<tr>
			<td>Plate photometer - Infinite M200</td>
			<td>Tecan</td>
			<td>NA</td>
			<td>Infinite M200 is discontinued. A Infinite 200 PRO is available at Tecan. 
			Software: SW Magellan v.6.3</td>
		</tr>
		<tr>
			<td>Potassium chloride</td>
			<td>Merck</td>
			<td>P9333</td>
			<td>For SIF buffer</td>
		</tr>
		<tr>
			<td>Scissor</td>
			<td>Allgaier Instruments</td>
			<td>307-156-170</td>
			<td></td>
		</tr>
		<tr>
			<td>Small scissor</td>
			<td>Allgaier Instruments</td>
			<td>04-520-115</td>
			<td></td>
		</tr>
		<tr>
			<td>Sodium bicarbonate</td>
			<td>Merck</td>
			<td>S6014</td>
			<td>For SIF buffer</td>
		</tr>
		<tr>
			<td>Sodium chloride</td>
			<td>Merck</td>
			<td>S9888</td>
			<td>For SIF buffer</td>
		</tr>
		<tr>
			<td>Sodium dihydrogen phosphate monohydrate</td>
			<td>Merck</td>
			<td>1.06346.1000</td>
			<td>For SIF buffer</td>
		</tr>
		<tr>
			<td>Sodium gluconate</td>
			<td>Merck</td>
			<td>S2054</td>
			<td>For SIF buffer</td>
		</tr>
		<tr>
			<td>Spatula</td>
			<td>Bochem Lab Supply</td>
			<td>3018</td>
			<td></td>
		</tr>
		<tr>
			<td>Spring scissor</td>
			<td>Fine Science Tools</td>
			<td>15024-10</td>
			<td></td>
		</tr>
		<tr>
			<td>Sucrose</td>
			<td>Merck</td>
			<td>84097</td>
			<td>For SIF buffer</td>
		</tr>
		<tr>
			<td>Supercinnamaldehyde</td>
			<td>Merck</td>
			<td>S3322</td>
			<td></td>
		</tr>
		<tr>
			<td>Tulle (fabrics)</td>
			<td>NA</td>
			<td>NA</td>
			<td>Bought in the local fabrics store</td>
		</tr>
	</tbody>
</table>

ex vivo release of calcitonin gene-related peptide from the trigeminovascular system in rodents

Calcitonin gene-related peptide (CGRP) was first discovered in the 1980s as a splice variant from the calcitonin gene. Since its discovery, its role in migraine pathophysiology has been well established, first by its potent vasodilator properties and subsequently by its presence and function as a neurotransmitter in the sensory trigeminovascular system. The migraine-provoking ability of CGRP gave support to the pharma industry to develop monoclonal antibodies and antagonists inhibiting the effect of CGRP. A new treatment paradigm has proven effective in the prophylactic treatment of migraine. One of the useful tools to further understand migraine mechanisms is the ex vivo model of CGRP release from the trigeminovascular system. It is a relatively simple method that can be used with various pharmacological tools to achieve know-how to further develop new effective migraine treatments. The present protocol describes a CGRP release model and the technique to quantify the effect of pharmacological agents on the amount of CGRP released from the trigeminovascular system in rodents. A procedure describing the experimental approach from euthanasia to the measurement of protein levels is provided. The essential isolation of the trigeminal ganglion and the trigeminal nucleus caudalis from both mice and rats and the preparation of rat dura mater are described in detail. Furthermore, representative results from both species (rats and mice) are presented. The technique is a key tool to investigate the molecular mechanisms involved in migraine pathophysiology by using various pharmacological compounds and genetically modified animals.

This technique is a tool to investigate the CGRP-related molecular mechanisms involved in migraine. It has the advantage of assessing CGRP release from different levels of the trigeminovascular system and can be applied both on wild-type and transgenic mice and rats in combination with various pharmacological compounds. Here, concentration-response and blocking experiments from rats and concentration-response results from wild-type and transgenic mice are presented.
The CGRP release method was used to study the effect of the KATP channel inhibitor glibenclamide on CGRP release from TG and dura mater in female spontaneous trigeminal allodynic (STA) rats. First, the optimal concentration of capsaicin was found using a concentration-response study design. Capsaicin exposure induced a significant CGRP release from dura mater and TG compared to the vehicle (Figure 5). In the dura mater, the maximum release of CGRP was found at 1 &#181;M of capsaicin, and in TG, the maximum CGRP release was found at 10 &#181;M of capsaicin (Figure 5A-B). Based on the concentration-response experiments, 1 &#181;M of capsaicin and 3 &#181;M of glibenclamide were used for blocking experiments. Glibenclamide showed no effect on basal CGRP release from dura mater (P = 0.441) and TG (P = 0.881) when analyzed with a one-way ANOVA51. Glibenclamide significantly reduced the capsaicin-induced CGRP release in dura mater by 40% (P = 0.031) and TG by 39% (P = 0.003) compared to capsaicin with the vehicle when analyzed with a one-way ANOVA (Figure 5C-D)51.
In mice, the protocol was used to examine the involvement of the transient receptor potential ankyrin 1 (TRPA1) ion channel in a GTN mouse model of migraine, where GTN-induced hypersensitivity was fully dependent on TRPA1 channels. The TRPA1 agonist supercinnamaldehyde (SCA) was found to release CGRP in a dose-dependent manner from the TG with 1 &#181;M, 10 &#181;M, and 100 &#181;M of SCA, resulting in 9% (P = 0.23), 51% (P = 0.011), and 69% (P = 0.0097) increased release of CGRP compared to vehicle, respectively when analyzed with two-way ANOVA. This release was absent in TG from Trpa1 null mice where exposure to 1 &#181;M, 10 &#181;M, and 100 &#181;M of SCA resulted in 11% (P &#62; 0.99), -13% (P &#62; 0.99) and 9% (P = 0.97) percent change in the release of CGRP compared to vehicle, respectively when analyzed with two-way ANOVA. The subsequent stimulation with 10 &#181;M of capsaicin (positive control) shows that all tissue samples could release CGRP (Figure 6)50.
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/63723/63723fig01.jpg" /> 
Figure 1: Step-by-step dissection of tissue from rats. The (A-T) details are provided in the protocol section (step 3.1). <a href="https://www.jove.com/files/ftp_upload/63723/63723fig01large.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/63723/63723fig02.jpg" /> 
Figure 2: Step-by-step dissection of tissue from mice. The (A-T) details are provided in the protocol section (step 3.2). <a href="https://www.jove.com/files/ftp_upload/63723/63723fig02large.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/63723/63723fig03.jpg" /> 
Figure 3: Washing and incubation of rodent tissue. (A-B) Freshly isolated tissue in plastic containers with SIF. The containers are covered with tulle to enable easy change of SIF. (C) Rat tissue - Two skull halves with dura mater covering the inner linings of the skull placed on a 6-well culture plate. The right and left trigeminal nucleus caudalis in separate microcentrifuge tube lids (top row of lids). The two trigeminal ganglia in individual microcentrifuge tube lids (bottom row of lids). (D) Mouse tissue - The trigeminal nucleus caudalis-containing part of the brainstem in a separate microcentrifuge tube lid (top). Two mouse trigeminal ganglia are in one microcentrifuge tube lid (bottom). <a href="https://www.jove.com/files/ftp_upload/63723/63723fig03large.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/63723/63723fig04.jpg" /> 
Figure 4: Study designs for CGRP release. Three different protocols for performing CGRP release experiments. Drugs are to be diluted in synthetic interstitial fluid (SIF). (A) Single stimulation. (B) Concentration-response stimulation. (C) Inhibition of a stimulation. <a href="https://www.jove.com/files/ftp_upload/63723/63723fig04large.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/63723/63723fig05.jpg" /> 
Figure 5: Glibenclamide inhibits capsaicin-induced CGRP release from rat dura mater and trigeminal ganglion. CGRP levels from trigeminal ganglion and dura mater isolated from female spontaneous trigeminal allodynic (STA) rats (215-318 g) originally sourced from the Thomas Jefferson University52 were measured with commercial human CGRP EIA kits. (A-B) CGRP release from (A) dura mater and (B) trigeminal ganglion after increasing concentrations of capsaicin (10 nM, 100 nM, 1 &#181;M and 10 &#181;M) (n = 4). Data are presented as individual points and were analyzed with two-way ANOVA. ****p &#60; 0.0001 (C-D) Levels of CGRP released from (C) dura mater and (D) trigeminal ganglion after 10 min exposure to vehicle, 3 &#181;M glibenclamide (glib), 1 &#181;M capsaicin and 1 &#181;M capsaicin + 3 &#181;M glib (n = 6-11). Data are presented as individual points and mean values and were analyzed with one-way ANOVA. *compared to vehicle. #capsaicin compared to capsaicin + glib. #P &#60; 0.05, **&#160;and&#160;##P &#60; 0.01, ****P &#60; 0.0001. The analyses were followed with Bonferroni&#39;s multiple comparison test. A significant level of &#945; = 0.05 was used for all the tests. This figure has been modified from Christensen et al.51. <a href="https://www.jove.com/files/ftp_upload/63723/63723fig05large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 6" class="xfigimg" src="/files/ftp_upload/63723/63723fig06.jpg" /> 
Figure 6: SCA exposure results in TRPA1 dependent CGRP release from mouse trigeminal ganglion. CGRP levels released from trigeminal ganglion isolated from WT and Trpa1 null (Trpa1tm1/Dpc)53 male mice (8-10 weeks) were measured with rat CGRP EIA kits after exposure to supercinnamaldehyde (SCA) at 1 &#181;M, 10 &#181;M, and 100 &#181;M and capsaicin at 10 &#181;M as a positive control (n = 6). Data from each mouse is presented as individual points, and bars indicate the mean values. Statistics: Comparison between SCA and vehicle at each concentration and between basal and positive control was performed with a two-way repeated ANOVA. A significant level of &#945; = 0.05. *P &#60; 0.05, **P &#60; 0.01. This figure has been modified from Christensen et al.50. <a href="https://www.jove.com/files/ftp_upload/63723/63723fig06large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

Watch this Scientific Journal Video about Ex Vivo Release of Calcitonin Gene-Related Peptide from the Trigeminovascular System in Rodents at JoVE.com

Ex Vivo Release of Calcitonin Gene-Related Peptide from the Trigeminovascular System in Rodents

The present protocol describes the ex vivo calcitonin gene-related peptide (CGRP) release model and the strategy to quantify the effect of pharmacological agents on the amount of CGRP released from the trigeminovascular system in rodents.

Ex Vivo Release of Calcitonin Gene-Related Peptide from the Trigeminovascular System in Rodents

Calcitonin gene-related peptide (CGRP) was first discovered in the 1980s as a splice variant from the calcitonin gene. Since its discovery, its role in ...

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2.2K Views.  Rigshospitalet, University of Copenhagen. The present protocol describes the ex vivo calcitonin gene-related peptide (CGRP) release model and the strategy to quantify the effect of pharmacological agents on the amount of CGRP released from the trigeminovascular system in rodents.

Video: Ex Vivo Release of Calcitonin Gene-Related Peptide from the Trigeminovascular System in Rodents

Isolating Nasal Olfactory Stem Cells from Rodents or Humans

The olfactory mucosa, located in the nasal cavity, is in charge of detecting odours. It is also the only nervous tissue that is exposed to the external environment and easily accessible in every living individual. As a result, this tissue is unique for anyone aiming to identify molecular anomalies in the pathological brain or isolate adult stem cells for cell therapy. 
Molecular abnormalities in brain diseases are often studied using nervous tissue samples collected post-mortem. However, this material has numerous limitations. In contrast, the olfactory mucosa is readily accessible and can be biopsied safely without any loss of sense of smell1. Accordingly, the olfactory mucosa provides an "open window" in the adult human through which one can study developmental (e.g. autism, schizophrenia)2-4 or neurodegenerative (e.g. Parkinson, Alzheimer) diseases4,5. Olfactory mucosa can be used for either comparative molecular studies4,6 or in vitro experiments on neurogenesis3,7.
The olfactory epithelium is also a nervous tissue that produces new neurons every day to replace those that are damaged by pollution, bacterial of viral infections. This permanent neurogenesis is sustained by progenitors but also stem cells residing within both compartments of the mucosa, namely the neuroepithelium and the underlying lamina propria8-10. We recently developed a method to purify the adult stem cells located in the lamina propria and, after having demonstrated that they are closely related to bone marrow mesenchymal stem cells (BM-MSC), we named them olfactory ecto-mesenchymal stem cells (OE-MSC)11.
Interestingly, when compared to BM-MSCs, OE-MSCs display a high proliferation rate, an elevated clonogenicity and an inclination to differentiate into neural cells. We took advantage of these characteristics to perform studies dedicated to unveil new candidate genes in schizophrenia and Parkinson's disease4. We and others have also shown that OE-MSCs are promising candidates for cell therapy, after a spinal cord trauma12,13, a cochlear damage14 or in an animal models of Parkinson's disease15 or amnesia16.
In this study, we present methods to biopsy olfactory mucosa in rats and humans. After collection, the lamina propria is enzymatically separated from the epithelium and stem cells are purified using an enzymatic or a non-enzymatic method. Purified olfactory stem cells can then be either grown in large numbers and banked in liquid nitrogen or induced to form spheres or differentiated into neural cells. These stem cells can also be used for comparative omics (genomic, transcriptomic, epigenomic, proteomic) studies.

We describe here a method for biopsying olfactory mucosa from rat and human nasal cavities. These biopsies can be used for either identifying molecular anomalies in brain diseases or isolating multipotent adult stem cells that can be utilized for cell transplantation in animal models of brain trauma/disease.

The olfactory mucosa, located in the nasal cavity, is in charge of detecting odours. It is also the only nervous tissue that is exposed to the external ...

A Fully Automated and Highly Versatile System for Testing Multi-cognitive Functions and Recording Neuronal Activities in Rodents

We have developed a fully automated system for operant behavior testing and neuronal activity recording by which multiple cognitive brain functions can be investigated in a single task sequence. The unique feature of this system is a custom-made, acoustically transparent chamber that eliminates many of the issues associated with auditory cue control in most commercially available chambers. The ease with which operant devices can be added or replaced makes this system quite versatile, allowing for the implementation of a variety of auditory, visual, and olfactory behavioral tasks. Automation of the system allows fine temporal (10 ms) control and precise time-stamping of each event in a predesigned behavioral sequence. When combined with a multi-channel electrophysiology recording system, multiple cognitive brain functions, such as motivation, attention, decision-making, patience, and rewards, can be examined sequentially or independently.

In this report, we present a fully automated and highly versatile system capable of simultaneously testing multi-cognitive behaviors and recording neuronal activities for rodents.

We have developed a fully automated system for operant behavior testing and neuronal activity recording by which multiple cognitive brain functions can be ...

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

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

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

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

In vivo Optogenetic Stimulation of the Rodent Central Nervous System

The ability to probe defined neural circuits in awake, freely-moving animals with cell-type specificity, spatial precision, and high temporal resolution has been a long sought tool for neuroscientists in the systems-level search for the neural circuitry governing complex behavioral states. Optogenetics is a cutting-edge tool that is revolutionizing the field of neuroscience and represents one of the first systematic approaches to enable causal testing regarding the relation between neural signaling events and behavior. By combining optical and genetic approaches, neural signaling can be bi-directionally controlled through expression of light-sensitive ion channels (opsins) in mammalian cells. The current protocol describes delivery of specific wavelengths of light to opsin-expressing cells in deep brain structures of awake, freely-moving rodents for neural circuit modulation. Theoretical principles of light transmission as an experimental consideration are discussed in the context of performing in vivo optogenetic stimulation. The protocol details the design and construction of both simple and complex laser configurations and describes tethering strategies to permit simultaneous stimulation of multiple animals for high-throughput behavioral testing.

In vivo Optogenetic Stimulation of the Rodent Central Nervous System

Optogenetics has become a powerful tool for use in behavioral neuroscience experiments. This protocol offers a step-by-step guide to the design and set-up of laser systems, and provides a full protocol for carrying out multiple and simultaneous in vivo optogenetic stimulations compatible with most rodent behavioral testing paradigms.

The ability to probe defined neural circuits in awake, freely-moving animals with cell-type specificity, spatial precision, and high temporal resolution ...

Ex Vivo Preparations of the Intact Vomeronasal Organ and Accessory Olfactory Bulb

The mouse accessory olfactory system (AOS) is a specialized sensory pathway for detecting nonvolatile social odors, pheromones, and kairomones. The first neural circuit in the AOS pathway, called the accessory olfactory bulb (AOB), plays an important role in establishing sex-typical behaviors such as territorial aggression and mating. This small (&#60;1 mm3) circuit possesses the capacity to distinguish unique behavioral states, such as sex, strain, and stress from chemosensory cues in the secretions and excretions of conspecifics. While the compact organization of this system presents unique opportunities for recording from large portions of the circuit simultaneously, investigation of sensory processing in the AOB remains challenging, largely due to its experimentally disadvantageous location in the brain. Here, we demonstrate a multi-stage dissection that removes the intact AOB inside a single hemisphere of the anterior mouse skull, leaving connections to both the peripheral vomeronasal sensory neurons (VSNs) and local neuronal circuitry intact. The procedure exposes the AOB surface to direct visual inspection, facilitating electrophysiological and optical recordings from AOB circuit elements in the absence of anesthetics. Upon inserting a thin cannula into the vomeronasal organ (VNO), which houses the VSNs, one can directly expose the periphery to social odors and pheromones while recording downstream activity in the AOB. This procedure enables controlled inquiries into AOS information processing, which can shed light on mechanisms linking pheromone exposure to changes in behavior.

Ex Vivo Preparations of the Intact Vomeronasal Organ and Accessory Olfactory Bulb

The mouse accessory olfactory bulb (AOB) has been difficult to study in the context of sensory coding. Here, we demonstrate a dissection that produces an ex vivo preparation in which AOB neurons remain functionally connected to their peripheral inputs, facilitating research into information processing of mouse pheromones and kairomones.

The mouse accessory olfactory system (AOS) is a specialized sensory pathway for detecting nonvolatile social odors, pheromones, and kairomones. The first ...

Simultaneous ex vivo Functional Testing of Two Retinas by in vivo Electroretinogram System

An In vivo electroretinogram (ERG) signal is composed of several overlapping components originating from different retinal cell types, as well as noise from extra-retinal sources. Ex vivo ERG provides an efficient method to dissect the function of retinal cells directly from an intact isolated retina of animals or donor eyes. In addition, ex vivo ERG can be used to test the efficacy and safety of potential therapeutic agents on retina tissue from animals or humans. We show here how commercially available in vivo ERG systems can be used to conduct ex vivo ERG recordings from isolated mouse retinas. We combine the light stimulation, electronic and heating units of a standard in vivo system with custom-designed specimen holder, gravity-controlled perfusion system and electromagnetic noise shielding to record low-noise ex vivo ERG signals simultaneously from two retinas with the acquisition software included in commercial in vivo systems. Further, we demonstrate how to use this method in combination with pharmacological treatments that remove specific ERG components in order to dissect the function of certain retinal cell types.

Simultaneous ex vivo Functional Testing of Two Retinas by in vivo Electroretinogram System

Ex vivo ERG can be used to record electrical activity of retinal cells directly from isolated intact retinas of animals or humans. We demonstrate here how common in vivo ERG systems can be adapted for ex vivo ERG recordings in order to dissect the electrical activity of retinal cells.

An In vivo electroretinogram (ERG) signal is composed of several overlapping components originating from different retinal cell types, as well as noise ...

Observation of the Ciliary Movement of Choroid Plexus Epithelial Cells Ex Vivo

The choroid plexus is located in the ventricular wall of the brain, the main function of which is believed to be production of cerebrospinal fluid. Choroid plexus epithelial cells (CPECs) covering the surface of choroid plexus tissue harbor multiple unique cilia, but most of the functions of these cilia remain to be investigated. To uncover the function of CPEC cilia with particular reference to their motility, an ex vivo observation system was developed to monitor ciliary motility during embryonic, perinatal and postnatal periods. The choroid plexus was dissected out of the brain ventricle and observed under a video-enhanced contrast microscope equipped with differential interference contrast optics. Under this condition, a simple and quantitative method was developed to analyze the motile profiles of CPEC cilia for several hours ex vivo. Next, the morphological changes of cilia during development were observed by scanning electron microscopy to elucidate the relationship between the morphological maturity of cilia and motility. Interestingly, this method could delineate changes in the number and length of cilia, which peaked at postnatal day (P) 2, while the beating frequency reached a maximum at P10, followed by abrupt cessation at P14. These techniques will enable elucidation of the functions of cilia in various tissues. While related techniques have been published in a previous report1, the current study focuses on detailed techniques to observe the motility and morphology of CPEC cilia ex vivo.

Observation of the Ciliary Movement of Choroid Plexus Epithelial Cells Ex Vivo

In this study, a detailed light microscopic technique was optimized for real-time observation and analysis of the motion of CPEC cilia ex vivo together with an electron microscopic method for ultrastructural analysis.

The choroid plexus is located in the ventricular wall of the brain, the main function of which is believed to be production of cerebrospinal fluid. ...

Ex Vivo Optogenetic Dissection of Fear Circuits in Brain Slices

Optogenetic approaches are now widely used to study the function of neural populations and circuits by combining targeted expression of light-activated proteins and subsequent manipulation of neural activity by light. Channelrhodopsins (ChRs) are light-gated cation-channels and when fused to a fluorescent protein their expression allows for visualization and concurrent activation of specific cell types and their axonal projections in defined areas of the brain. Via stereotactic injection of viral vectors, ChR fusion proteins can be constitutively or conditionally expressed in specific cells of a defined brain region, and their axonal projections can subsequently be studied anatomically and functionally via ex vivo optogenetic activation in brain slices. This is of particular importance when aiming to understand synaptic properties of connections that could not be addressed with conventional electrical stimulation approaches, or in identifying novel afferent and efferent connectivity that was previously poorly understood. Here, a few examples illustrate how this technique can be applied to investigate these questions to elucidating fear-related circuits in the amygdala. The amygdala is a key region for acquisition and expression of fear, and storage of fear and emotional memories. Many lines of evidence suggest that the medial prefrontal cortex (mPFC) participates in different aspects of fear acquisition and extinction, but its precise connectivity with the amygdala is just starting to be understood. First, it is shown how ex vivo optogenetic activation can be used to study aspects of synaptic communication between mPFC afferents and target cells in the basolateral amygdala (BLA). Furthermore, it is illustrated how this ex vivo optogenetic approach can be applied to assess novel connectivity patterns using a group of GABAergic neurons in the amygdala, the paracapsular intercalated cell cluster (mpITC), as an example.

Ex Vivo Optogenetic Dissection of Fear Circuits in Brain Slices

Optogenetic approaches are widely used to manipulate neural activity and assess the consequences for brain function. Here, a technique is outlined that&#160;upon in vivo expression of the optical activator Channelrhodopsin, allows for ex vivo analysis of synaptic properties of specific long range and local neural connections in fear-related circuits.

Optogenetic approaches are now widely used to study the function of neural populations and circuits by combining targeted expression of light-activated ...

The Power of Interstimulus Interval for the Assessment of Temporal Processing in Rodents

Temporal processing deficits have been implicated as a potential elemental dimension of higher-level cognitive processes, commonly observed in neurocognitive disorders. Despite the popularization of prepulse inhibition (PPI) in recent years, many current protocols promote using a percent of control measure, thereby precluding the assessment of temporal processing. The present study used cross-modal PPI and gap prepulse inhibition (gap-PPI) to demonstrate the benefits of employing a range of interstimulus intervals (ISIs) to delineate effects of sensory modality, psychostimulant exposure, and age. Assessment of sensory modality, psychostimulant exposure, and age reveals the utility of an approach varying the interstimulus interval (ISI) to establish the shape of the ISI function, including increases (sharper curve inflections) or decreases (flattening of the response amplitude curve) in startle amplitude. Additionally, shifts in peak response inhibition, suggestive of a differential sensitivity to the manipulation of ISI, are often revealed. Thus, the systematic manipulation of ISI affords a critical opportunity to evaluate temporal processing, which may reveal the underlying neural mechanisms involved in neurocognitive disorders.

Temporal processing, a preattentive process, may underlie deficits in higher-level cognitive processes, including attention, commonly observed in neurocognitive disorders. Using prepulse inhibition as an exemplar paradigm, we present a protocol for manipulating interstimulus interval (ISI) to establish the shape of the ISI function to provide an assessment of temporal processing.

Temporal processing deficits have been implicated as a potential elemental dimension of higher-level cognitive processes, commonly observed in ...

Visualizing the Calcitonin Gene-Related Peptide Immunoreactive Innervation of the Rat Cranial Dura Mater with Immunofluorescence and Neural Tracing

The aim of this study was to examine the distribution and origin of the calcitonin gene-related peptide (CGRP)-immunoreactive sensory nerve fibers of the cranial dura mater using immunofluorescence, three-dimensional (3D) reconstruction and retrograde tracing technique. Here, the nerve fibers and blood vessels were stained using immunofluorescence and histochemistry techniques with CGRP and fluorescent phalloidin, respectively. The spatial correlation of dural CGRP-immuoreactive nerve fibers and blood vessels were demonstrated by 3D reconstruction. Meanwhile, the origin of the CGRP-immunoreactive nerve fibers were detected by neural tracing technique with fluorogold (FG) from the area around middle meningeal artery (MMA) in the cranial dura mater to the trigeminal ganglion (TG) and cervical (C) dorsal root ganglia (DRGs). In addition, the chemical characteristics of FG-labeled neurons in the TG and DRGs were also examined together with CGRP using double immunofluorescences. Taking advantage of the transparent whole-mount sample and 3D reconstruction, it was shown that CGRP-immunoreactive nerve fibers and phalloidin-labeled arterioles run together or separately forming a dural neurovascular network in a 3D view, while the FG-labeled neurons were found in the ophthalmic, maxillary, and mandibular branches of TG, as well as the C2-3 DRGs ipsilateral to the side of tracer application in which some of FG-labeled neurons presented with CGRP-immunoreactive expression. With these approaches, we demonstrated the distributional characteristics of CGRP-immunoreactive nerve fibers around the blood vessels in the cranial dura mater, as well as the origin of these nerve fibers from TG and DRGs. From the perspective of methodology, it may provide a valuable reference for understanding the complicated neurovascular structure of the cranial dura mater under the physiological or pathological condition.

Here we present a protocol to visualize spatial correlation of calcitonin gene-related peptide (CGRP)-immunoreactive nerve fibers and blood vessels in the cranial dura mater using immunofluorescence and fluorescent histochemistry with CGRP and phalloidin, respectively. In addition, the origin of these nerve fibers was retrograde traced with a fluorescent neural tracer.

The aim of this study was to examine the distribution and origin of the calcitonin gene-related peptide (CGRP)-immunoreactive sensory nerve fibers of the ...