This study was supported by the National Institutes of Health (NS104200 and NS072204 to GD).

All procedures were conducted with prior approval of the institutional Animal Care and Use Committee at the University of Texas at Dallas. ICR (CD-1) (30-35 g) and C57/BL6 (25-30 g) mice aged 6-8 weeks were used in this study.
1. Dural infuser
<ol>
	<li>Create the mouse infusers/injectors by modifying a commercially available internal cannula and infuser for unilateral injections with a non-metallic fused silica plastic cap that is adjustable and inserts into/extends below a 28 G guide cannula with inner diameter (I.D.) of 0.18 mm and an outer diameter (O.D.) of 0.35 mm (Figure 1A).</li>
	<li>Use a caliper or other measuring device to adjust the fused silica plastic cap on the infuser to a length of 0.6 mm; measured from the tip of the infuser to the edge of the silica plastic cap.
	<ol>
		<li>Take caution not to bend or dull the infuser when adjusting the plastic cap.</li>
		<li>For other mouse strains that have not been previously used for dural injections, determine the optimal infuser length by setting the length to 0.6 mm and conducting pilot injections with ink or dye, adjusting the infuser length until it is observed that the dye is only in the dura mater and not on the brain or skull.</li>
	</ol>
	</li>
	<li>Attach the long end (or the end that was not measured to be 0.6 mm) of the adjusted infuser to plastic tubing (pump tubing, 2-stop, I.D. 0.19 mm, length 406 mm).
	<ol>
		<li>Cut the tubing to a minimum length of 8 in to ensure there is sufficient line to hold a volume of 5 &#181;L.</li>
		<li>Make sure the tubing covers the metal part and the top of the plastic stopper located on the infuser. This will help prevent air bubbles from accumulating in the line.</li>
	</ol>
	</li>
	<li>Attach the other end of the tubing to a 10 &#181;L glass microsyringe (gas-tight; cemented needle; 21 G with a 10 mm projection), again, making sure to have a tight seal over the metal part of the syringe (Figure 1A).</li>
	<li>Once the line is connected, backfill the syringe with 5 &#181;L of phosphate-buffered saline (PBS), synthetic interstitial fluid (SIF), or other vehicles of choice to prevent air bubbles from forming.
	<ol>
		<li>If air bubbles are observed in the line, flood the line with the vehicle until the bubbles have dissipated. 
		NOTE: It may help to fill the syringe with the vehicle prior to attaching it to the line and then push the fluid through the line once connected.</li>
	</ol>
	</li>
	<li>After the line is backfilled with 5 &#181;L of the vehicle and is working efficiently, load 5 &#181;L of the drug/solution into Hamilton syringe (ink or dye may be used as an alternative to the drug/solution if learning or practicing this technique).
	<ol>
		<li>Ensure that all vehicle solutions administered onto the dura are maintained at pH 7.4 and measured to an osmolality of 310. This reduces the potential activation of acid-sensing ion channels and other osmosensitive channels within the dura.</li>
	</ol>
	</li>
	<li>The maximum volume tested in mice that did not cause leaking into the brain is roughly 10 &#181;L. The behavioral effects after injections with this volume have not been tested. For this reason, administer only 5 &#181;L of the solution onto the dura. 
	&#8203;NOTE: These observations are based on the mouse strains/ages/weights of 6-8 week old CD1/ICR mice.</li>
</ol>
2. Dural injections
<ol>
	<li>Once the syringe is prepared and the drug is loaded, position a mouse flat onto its abdomen and anesthetize it under brief 3% isoflurane with an oxygen flow rate of 0.5-1 L/min via nosecone.
	<ol>
		<li>After the mouse no longer displays a pinch reflex, adjust the anesthesia and sustain it at a 1.5% isoflurane.</li>
	</ol>
	</li>
	<li>Once anesthetized, apply sterile opthalmic ointment to the eyes and shave the head of the animal, then disinfect the skin with povidone-iodine and ethanol. Following this, get&#160;in a position conducive to a successful injection.</li>
	<li>Use one hand to steady the head of the animal and hold the infuser with the other hand.</li>
	<li>Carefully probe and locate the junction of the sagittal and lambdoidal sutures on the skull of the mouse (Figure 1B, C).
	<ol>
		<li>To locate this discreet junction through the skin, use the topographical features of the skull and gently probe the general location of the junction with the infuser.</li>
		<li>Verify the position of the junction by re-positioning the infuser along the skull and feeling for the exact location.</li>
	</ol>
	</li>
	<li>Once the suture is located and the infuser is in place, very slowly and gently wiggle the infuser back and forth until it pierces through the skin and drops down into the junction all the way up to the plastic stopper. 
	NOTE: Ensure to insert the entire 0.6 mm tip of the infuser into the junction.</li>
	<li>To verify the accuracy, use ink or dye as an injection solution and euthanize and decapitate the mouse.
	<ol>
		<li>Remove the skull cap to visualize the dye within the dura mater (Figure 1C). 
		NOTE: Dye should not be observed on the brain or exterior of the skull. Likewise, mice in any experiment should be checked post-mortem to verify the accuracy of the injection as well as to ensure the integrity of the dura mater was undamaged.</li>
	</ol>
	</li>
	<li>Following the injection, remove the mouse from anesthesia, wait for it to regain consciousness and then return to its cage or place it into a testing chamber to begin the desired assays. 
	&#8203;NOTE: Allow the mouse to recover from anesthesia for a minimum of 30 min prior to doing any behavioral experiments.</li>
</ol>
3. Periorbital von Frey
<ol>
	<li>Begin the study with a cohort of approximately 16-20 mice.</li>
	<li>One the day prior to habituation, handle each mouse for at least 5 min.</li>
	<li>Approximately 24 h after handling, habituate the mice to the testing room conditions and the von Frey testing apparatus (Figure 2A). 
	NOTE: The acrylic testing apparatus consists of individual compartments with lids that are approximately 3 in x 3.5 in x 5 in (W x H x D) and are supported by aluminum stands connected via 0.25 in 19 G square galvanized steel mesh wire.
	<ol>
		<li>Place the mice inside a horizontally placed 4-oz white paper cup that is odorless and does not contain polythene or paraffin wax. 
		NOTE: These types of cups are preferred because it reduces gastrointestinal upset in the mice if ingested (Figure 2B).</li>
	</ol>
	</li>
	<li>While the animals are in their respective chambers, place a pellet of the normal chow diet in the individual chamber of each mouse to calm the animals and avoids any unnecessary stress to the animals. Do this for 3 days prior to any von Frey behavior testing.
	<ol>
		<li>Ensure that every time the mice are in the chamber that there is access to food.</li>
		<li>Number and assign each animal to the same space in the testing rack. Place the mouse in the same cup every day of the testing period to ensure that each animal becomes acclimated to its testing environment. 
		NOTE: Mice will gnaw on the cups and subsequently destroy the cups. If this should happen, replace the cup and label it with the corresponding mouse number.</li>
	</ol>
	</li>
	<li>Following the initial 3 days of habituation, place the mice in their individual chambers.
	<ol>
		<li>Allow the animals to acclimate to the testing room and chambers for at least 1 h prior to any von Frey testing to allow the mice to calm down and subsequently are easier to test.</li>
	</ol>
	</li>
	<li>After acclimation to the room on the testing day, remove one mouse while still in its cup from its respective chamber.
	<ol>
		<li>Maintain the cup in the horizontal position so that the mouse is on both forepaws and hind paws to help keep their weight evenly distributed. 
		NOTE: Unequal weight distribution may alter animal&#39;s responses and even prevent animals from responding.</li>
	</ol>
	</li>
	<li>Place the cup with the mouse inside on the table below the testing rack on the absorbent pad.</li>
	<li>For periorbital von Frey testing, place the 0.07 g von Frey filament directly in the center of the face and between the eyes.</li>
	<li>Apply enough pressure on the filament to cause the von Frey hair to bend into a &#34;C&#34; shaped formation.
	<ol>
		<li>Maintain contact with the region at least 3 s but no more than 5 s or until the mouse withdraws its head and swipes at the filament with its paw. 
		NOTE: If the filament slips or more than the tip of the filament touches the animal during testing, any responses should not be counted. These responses may be in response to the brush which are activated by different mechanoreceptors and therefore may not reflect accurate results.</li>
		<li>Apply von Frey filaments according to the Dixon &#34;up-down&#34; method22,23.
		<ol>
			<li>Initially, apply the von Frey filament which has a weight of 0.07 g. The lowest possible filament and the highest tested filament in this study are filaments with weights of 0.008 g and 0.6 g, respectively.</li>
			<li>Use the filaments of weight 0.008 g, 0.02 g, 0.04 g, 0.07 g, 0.16 g, 0.4 g, and 0.6 g to perform this assay.</li>
			<li>In this method, if an animal does not exhibit a response to the filament, apply the filament of the next higher gram weight.</li>
			<li>If the mouse does respond to a filament, consider that mouse responsive to that filament. If this is the case, apply the filament of the next lower gram weight.</li>
			<li>Repeat this pattern until the animal is tested 4 times after the initial response or the animal is determined to be unresponsive to any filaments tested in the assay. 
			&#8203;NOTE: Refrain from applying any additional pressure stemming from the arm or wrist. A scale may be used to practice the application of the filament.</li>
		</ol>
		</li>
	</ol>
	</li>
</ol>
4. Testing for baseline withdrawal thresholds
<ol>
	<li>Prior to inclusion in an experiment, ensure that the mice reach a baseline withdrawal threshold between 0.5-0.6 g.
	<ol>
		<li>A mouse reaches baseline if they fail to respond to any filament tested in the series mentioned in step 3.9.2.2 (0.07 g, 0.16 g, 0.4 g, and 0.6 g).</li>
	</ol>
	</li>
	<li>Test mice daily when establishing baseline withdrawal thresholds.
	<ol>
		<li>Testing allows animals to acclimate to the testing conditions and the pressure of the von Frey filaments.</li>
		<li>If the mice are still extremely hypersensitive after the third day of testing, try waiting 1 or 2 days before testing again. 
		NOTE: Too much time between testing days may result in the animal failing to adjust to the weight of the filament on their periorbital region, thus not reaching the targeted withdrawal threshold.</li>
	</ol>
	</li>
	<li>Test mice for approximately 7 days before determining which animals do not meet the inclusion criteria for an experiment. 
	&#8203;NOTE: Approximately 70% of mice will reach the targeted baseline level.
	<ol>
		<li>Prior to dural stimulation, analyze the baseline data to exclude any mouse that has not reached a baseline value of 0.5-0.6 grams or higher.</li>
		<li>After exclusion, randomly allocate each remaining mouse to a testing group. Achieve this by drawing out of a cup or writing a script on a spreadsheet to randomize numbers to a group.</li>
	</ol>
	</li>
</ol>
5. Analysis of von Frey results
<ol>
	<li>Once the series of responses have been obtained, determine the delta, k value, the 50% threshold, and the withdrawal threshold in grams according to previously published methods24.</li>
	<li>Calculate the withdrawal threshold using this formula WT = 10(x*F+B), where WT= withdrawal threshold, F = paw withdrawal threshold calculated via the Chaplan method, and B = linear regression of log (bending force) = x*Filament number + B.</li>
	<li>Plot the data as either 50% withdrawal threshold or average withdrawal threshold in grams.</li>
</ol>

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The authors have nothing to disclose.

Maladaptive changes in the local nociceptive system in the dura are considered a key contributor to the headache phase of migraine attacks despite a lack of tissue injury25,26. Here the study presents a method whereby minimally invasive stimulation of the dura can induce facial tactile hypersensitivity. Elucidating the mechanisms and events involved in dural nociceptor activation without causing damage to the cranium and tissues may more accurately reflect migrai...

Migraine pain remains a major public health issue worldwide. The World Health Organization ranks it as the sixth-most prevalent disease in the world, afflicting just under 15% of the Earth&#39;s population1 and causing a substantial socioeconomic burden on society2,3. Treatment options and their efficacy have been suboptimal and only provide symptomatic relief and do not significantly modify pathophysiological events that underly migraine occurrence4,5. The lack of treatment success is likely due to migraine being a multifactorial disorder whose pathology is poorly understood, leading to a limited number of therapeutic targets. Migraine is also challenging to fully capture in animal models, especially given that migraine diagnosis is made based on verbal communication with patients who describe their experience with migraine hallmarks such as aura, headache, photophobia, and allodynia. Notwithstanding, it is important to note that recent advances in migraine treatments are currently outperforming treatments for many neurological conditions that have been well validated by preclinical models. For instance, monoclonal antibodies and small molecules that target calcitonin gene-related peptide, or its receptor have been very successful in improving the quality of life of migraine sufferers and can potentially transform the clinical management of migraine. While there has been advancement in understanding this disorder, there continues to be much yet to be elucidated.
Based on preclinical animal models and human studies, it is widely accepted that migraine headaches are initiated by aberrant activation of nociceptive fibers within the meninges that signal through the trigeminal and upper cervical dorsal-root ganglia6,7,8,9,10. Despite this theory, many studies still use systemic administration of drugs to understand underlying contributing mechanisms in migraine. While systemic dosing of drugs has substantially strengthened our understanding, these findings do not directly assess whether local actions within the target tissue of interest play a role in migraine. Conversely, several studies have taken an approach to stimulate the dura; however, these experiments require cannula implantation via an invasive craniotomy and extended recovery times11,12. Because of these limitations, we developed a minimally invasive approach to locally stimulate the dura where the lack of a craniotomy eliminates post-surgical recovery and allows for immediate testing in awake animals12,13,14. These injections are performed under light isoflurane anesthesia and administered at the junction of the sagittal and lambdoid sutures in mice.
Several approaches have been developed to evaluate nociceptive behavioral responses in rodents15. Cutaneous allodynia has been reported in approximately 80% of migraine sufferers16,17 and represents a potential translational endpoint for use in rodents. In preclinical models, the application of von Frey filaments to the plantar region of the rodent paw has been used to assess pain behaviors in preclinical migraine models. The primary limitation of this approach is that it does not test the cephalic region. Facial grimace scoring has been used to capture pain behaviors in rodents by analyzing facial expressions after induction of pain stimuli18,19. However, its limitations include only capturing responses to acute stimuli and not chronic orofacial pain conditions. Facial grooming and decreased rearing are also considered outputs of behavioral responses in preclinical models of migraine20,21. Limitations of the former include difficulty in differentiating pain responses from normal routine grooming and other sensations such as itch. In the case of the latter, rearing behaviors typically decrease quickly after the introduction of rodents to novel environments. Although each of these behavioral endpoints is valuable in the understanding of various mechanisms that contribute to pain conditions, there is a critical need for preclinical models of pain disorders such as migraine to include endpoints that specifically capture cephalic hypersensitivity responses. Assessing tactile hypersensitivity of the periorbital skin following dural stimulation is a method that may provide better insight into mechanisms contributing to migraines where sensory symptoms are predominantly cephalic in nature. Here, we describe a method to administer substances onto the mouse dura as a preclinical model of migraine. Following dural application, we also present a detailed method for testing periorbital tactile hypersensitivity using calibrated von Frey filaments applied in the Dixon up-down method.

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			<td>Custom</td>
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			<td>The galvanized steel chicken wire dimensions are 0.25 in. x 19-gauge</td>
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			<td>Testing Rack with individual&#160; Chambers</td>
			<td>Custom</td>
			<td></td>
			<td>Each chamber should have a division between each mouse and lids to contain the mouse. The chambers should also be large enough to hold a 4 oz. paper cup.</td>
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			<td>von Frey Filaments</td>
			<td>Touch test/Stoelting</td>
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dural stimulation and periorbital von frey testing in mice as a preclinical model of headache

The cranial meninges, comprised of the dura mater, arachnoid, and pia mater, are thought to primarily serve structural functions for the nervous system. For example, they protect the brain from the skull and anchor/organize the vascular and neuronal supply of the cortex. However, the meninges are also implicated in nervous system disorders such as migraine, where the pain experienced during a migraine is attributed to local sterile inflammation and subsequent activation of local nociceptive afferents. Of the layers in the meninges, the dura mater is of particular interest in the pathophysiology of migraines. It is highly vascularized, harbors local nociceptive neurons, and is home to a diverse array of resident cells such as immune cells. Subtle changes in the local meningeal microenvironment may lead to activation and sensitization of dural perivascular nociceptors, thus leading to migraine pain. Studies have sought to address how dural afferents become activated/sensitized by using either in vivo electrophysiology, imaging techniques, or behavioral models, but these commonly require very invasive surgeries. This protocol presents a method for comparatively non-invasive application of compounds on the dura mater in mice and a suitable method for measuring headache-like tactile sensitivity using periorbital von Frey testing following dural stimulation. This method maintains the integrity of the dura and skull and reduces confounding effects from invasive techniques by injecting substances through a 0.65 mm modified cannula at the junction of unfused sagittal and lambdoid sutures. This preclinical model will allow researchers to investigate a wide range of dural stimuli and their role in the pathological progression of migraine, such as nociceptor activation, immune cell activation, vascular changes, and pain behaviors, all while maintaining injury-free conditions to the skull and meninges.

This injection method is used to administer stimuli onto the dura of mice so that subsequent behavioral testing may occur. The most common behavioral output measured with this model is cutaneous facial hypersensitivity assessed via von Frey12,13,14. Here we show how this model can be used to assess potential sex-specific contributions to migraine pathology (Figure 3).
Thi...

Watch this Scientific Journal Video about Dural Stimulation and Periorbital von Frey Testing in Mice As a Preclinical Model of Headache at JoVE.com

Dural Stimulation and Periorbital von Frey Testing in Mice As a Preclinical Model of Headache

The most notable symptom of migraine is severe head pain, and it is hypothesized that this is mediated by sensory neurons innervating the meninges. Here, we present a method to locally apply substances to the dura in a minimally invasive manner while using facial hypersensitivity as an output.

The cranial meninges, comprised of the dura mater, arachnoid, and pia mater, are thought to primarily serve structural functions for the nervous system.  ...

dural-stimulation-periorbital-von-frey-testing-mice-as-preclinical

Research

JoVE Journal

Neuroscience

7.3K Views.  University of Texas at Dallas. The most notable symptom of migraine is severe head pain, and it is hypothesized that this is mediated by sensory neurons innervating the meninges. Here, we present a method to locally apply substances to the dura in a minimally invasive manner while using facial hypersensitivity as an output. 

Video: Dural Stimulation and Periorbital von Frey Testing in Mice As a Preclinical Model of Headache

Combining Transcranial Magnetic Stimulation and fMRI to Examine the Default Mode Network

The default mode network is a group of brain regions that are active when an individual is not focused on the outside world and the brain is at "wakeful rest."1,2,3 It is thought the default mode network corresponds to self-referential or "internal mentation".2,3
It has been hypothesized that, in humans, activity within the default mode network is correlated with certain pathologies (for instance, hyper-activation has been linked to schizophrenia 4,5,6 and autism spectrum disorders 7 whilst hypo-activation of the network has been linked to Alzheimer's and other neurodegenerative diseases 8). As such, noninvasive modulation of this network may represent a potential therapeutic intervention for a number of neurological and psychiatric pathologies linked to abnormal network activation. One possible tool to effect this modulation is Transcranial Magnetic Stimulation: a non-invasive neurostimulatory and neuromodulatory technique that can transiently or lastingly modulate cortical excitability (either increasing or decreasing it) via the application of localized magnetic field pulses.9
In order to explore the default mode network's propensity towards and tolerance of modulation, we will be combining TMS (to the left inferior parietal lobe) with functional magnetic resonance imaging (fMRI). Through this article, we will examine the protocol and considerations necessary to successfully combine these two neuroscientific tools.

In this article, we examine the methodology and considerations relevant to the combination of TMS and fMRI to examine the effects of brain stimulation on the default network.

The default mode network is a group of brain regions that are active when an individual is not focused on the outside world and the brain is at "wakeful ...

TMS: Using the Theta-Burst Protocol to Explore Mechanism of Plasticity in Individuals with Fragile X Syndrome and Autism

Fragile X Syndrome (FXS), also known as Martin-Bell Syndrome, is a genetic abnormality found on the X chromosome.1,2 Individuals suffering from FXS display abnormalities in the expression of FMR1 - a protein required for typical, healthy neural development.3 Recent data has suggested that the loss of this protein can cause the cortex to be hyperexcitable thereby affecting overall patterns of neural plasticity.4,5
In addition, Fragile X shows a strong comorbidity with autism: in fact, 30% of children with FXS are diagnosed with autism, and 2 - 5% of autistic children suffer from FXS.6
Transcranial Magnetic Stimulation (a non-invasive neurostimulatory and neuromodulatory technique that can transiently or lastingly modulate cortical excitability via the application of localized magnetic field pulses 7,8) represents a unique method of exploring plasticity and the manifestations of FXS within affected individuals. More specifically, Theta-Burst Stimulation (TBS), a specific stimulatory protocol shown to modulate cortical plasticity for a duration up to 30 minutes after stimulation cessation in healthy populations, has already proven an efficacious tool in the exploration of abnormal plasticity.9,10 
Recent studies have shown the effects of TBS last considerably longer in individuals on the autistic spectrum - up to 90 minutes.11 This extended effect-duration suggests an underlying abnormality in the brain's natural plasticity state in autistic individuals - similar to the hyperexcitability induced by Fragile X Syndrome.
In this experiment, utilizing single-pulse motor-evoked potentials (MEPs) as our benchmark, we will explore the effects of both intermittent and continuous TBS on cortical plasticity in individuals suffering from FXS and individuals on the Autistic Spectrum.

In this article, we examine the effects of Theta-Burst TMS stimulation on cortical plasticity in individuals suffering from Fragile X syndrome and individuals on the autistic spectrum.

Fragile X Syndrome (FXS), also known as Martin-Bell Syndrome, is a genetic abnormality found on the X chromosome.1,2 Individuals suffering from FXS ...

The NeuroStar TMS Device: Conducting the FDA Approved Protocol for Treatment of Depression

The Neuronetics NeuroStar Transcranial Magnetic Stimulation (TMS) System is a class II medical device that produces brief duration, pulsed magnetic fields. These rapidly alternating fields induce electrical currents within localized, targeted regions of the cortex which are associated with various physiological and functional brain changes.1,2,3
In 2007, O'Reardon et al., utilizing the NeuroStar device, published the results of an industry-sponsored, multisite, randomized, sham-stimulation controlled clinical trial in which 301 patients with major depression, who had previously failed to respond to at least one adequate antidepressant treatment trial, underwent either active or sham TMS over the left dorsolateral prefrontal cortex (DLPFC). The patients, who were medication-free at the time of the study, received TMS five times per week over 4-6 weeks.4
The results demonstrated that a sub-population of patients (those who were relatively less resistant to medication, having failed not more than two good pharmacologic trials) showed a statistically significant improvement on the Montgomery-Asberg Depression Scale (MADRS), the Hamilton Depression Rating Scale (HAMD), and various other outcome measures. In October 2008, supported by these and other similar results5,6,7, Neuronetics obtained the first and only Food and Drug Administration (FDA) approval for the clinical treatment of a specific form of medication-refractory depression using a TMS Therapy device (FDA approval K061053).
In this paper, we will explore the specified FDA approved NeuroStar depression treatment protocol (to be administered only under prescription and by a licensed medical profession in either an in- or outpatient setting).

In this article, we examine the methodology and considerations relevant to the FDA approved depression treatment protocol using the Neuronetics NeuroStar TMS device.

The Neuronetics NeuroStar Transcranial Magnetic Stimulation (TMS) System is a class II medical device that produces brief duration, pulsed magnetic ...

Electrode Positioning and Montage in Transcranial Direct Current Stimulation

Transcranial direct current stimulation (tDCS) is a technique that has been intensively investigated in the past decade as this method offers a non-invasive and safe alternative to change cortical excitability2. The effects of one session of tDCS can last for several minutes, and its effects depend on polarity of stimulation, such as that cathodal stimulation induces a decrease in cortical excitability, and anodal stimulation induces an increase in cortical excitability that may last beyond the duration of stimulation6. These effects have been explored in cognitive neuroscience and also clinically in a variety of neuropsychiatric disorders &#x2013; especially when applied over several consecutive sessions4. One area that has been attracting attention of neuroscientists and clinicians is the use of tDCS for modulation of pain-related neural networks3,5. Modulation of two main cortical areas in pain research has been explored: primary motor cortex and dorsolateral prefrontal cortex7. Due to the critical role of electrode montage, in this article, we show different alternatives for electrode placement for tDCS clinical trials on pain; discussing advantages and disadvantages of each method of stimulation.

Transcranial direct current stimulation (tDCS) is an established technique to modulate cortical excitability1,2. It has been used as an investigative tool in neuroscience due to its effects on cortical plasticity, easy operation, and safe profile. One area that tDCS has been showing encouraging results is pain alleviation 3-5.

Transcranial direct current stimulation (tDCS) is a technique that has been intensively investigated in the past decade as this method offers a ...

Measuring Changes in Tactile Sensitivity in the Hind Paw of Mice Using an Electronic von Frey Apparatus

Measuring inflammation-induced changes in thresholds of hind paw withdrawal from mechanical pressure is a useful technique to assess changes in pain perception in rodents. Withdrawal thresholds can be measured first at baseline and then following drug, venom, injury, allergen,&#160;or otherwise evoked inflammation by applying an accurate force on very specific areas of the skin. An electronic von Frey apparatus allows precise assessment of mouse hind paw withdrawal thresholds that are not limited by the available filament sizes in contrast to classical von Frey measurements. The ease and rapidity of measurements allow for incorporation of assessment of tactile sensitivity outcomes in diverse models of rapid-onset inflammatory and neuropathic pain as multiple measurements can be taken within a short time period. Experimental measurements for individual rodent subjects can be internally controlled against individual baseline responses and exclusion criteria easily established to standardize baseline responses within and across experimental groups. Thus, measurements using an electronic von Frey apparatus represent a useful modification of the well-established classical von Frey filament-based assays for rodent mechanical allodynia that may also be applied to other nonhuman mammalian models.

Electronic von Frey measurements are an effective and improved alternative to classical von Frey filaments, as they allow rapid and precise measurement of changes in rodent mechanical withdrawal thresholds to continuously applied pressure stimuli before and after an inflammatory event.

Measuring inflammation-induced changes in thresholds of hind paw withdrawal from mechanical pressure is a useful technique to assess changes in pain ...

A Simple and Inexpensive Method for Determining Cold Sensitivity and Adaptation in Mice

Cold hypersensitivity is a serious clinical problem, affecting a broad subset of patients and causing significant decreases in quality of life. The cold plantar assay allows the objective and inexpensive assessment of cold sensitivity in mice, and can quantify both analgesia and hypersensitivity. Mice are acclimated on a glass plate, and a compressed dry ice pellet is held against the glass surface underneath the hindpaw. The latency to withdrawal from the cooling glass is used as a measure of cold sensitivity.
Cold sensation is also important for survival in regions with seasonal temperature shifts, and in order to maintain sensitivity animals must be able to adjust their thermal response thresholds to match the ambient temperature. The Cold Plantar Assay (CPA) also allows the study of adaptation to changes in ambient temperature by testing the cold sensitivity of mice at temperatures ranging from 30 &#176;C to 5 &#176;C. Mice are acclimated as described above, but the glass plate is cooled to the desired starting temperature using aluminum boxes (or aluminum foil packets) filled with hot water, wet ice, or dry ice. The temperature of the plate is measured at the center using a filament T-type thermocouple probe. Once the plate has reached the desired starting temperature, the animals are tested as described above.
This assay allows testing of mice at temperatures ranging from innocuous to noxious. The CPA yields unambiguous and consistent behavioral responses in uninjured mice and can be used to quantify both hypersensitivity and analgesia. This protocol describes how to use the CPA to measure cold hypersensitivity, analgesia, and adaptation in mice.

The Cold Plantar Assay (CPA) measures cold responsiveness between 30 &#176;C and 5 &#176;C, and can also measure cold adaptation. This protocol describes how to use the CPA to measure cold hypersensitivity, analgesia, and adaptation in mice.

Cold hypersensitivity is a serious clinical problem, affecting a broad subset of patients and causing significant decreases in quality of life.  The cold ...

Transcranial Direct Current Stimulation (tDCS) in Mice

Transcranial direct current stimulation (tDCS) is a non-invasive neuromodulation technique proposed as an alternative or complementary treatment for several neuropsychiatric diseases. The biological effects of tDCS are not fully understood, which is in part explained due to the difficulty in obtaining human brain tissue. This protocol describes a tDCS mouse model that uses a chronically implanted electrode allowing the study of the long-lasting biological effects of tDCS. In this experimental model, tDCS changes the cortical gene expression and offers a prominent contribution to the understanding of the rationale for its therapeutic use.

Transcranial Direct Current Stimulation tDCS in Mice

Transcranial direct current stimulation (tDCS) is a therapeutic technique proposed to treat psychiatric diseases. An animal model is essential for understanding the specific biological alterations evoked by tDCS. This protocol describes a tDCS mouse model that uses a chronically implanted electrode.

Transcranial direct current stimulation (tDCS) is a non-invasive neuromodulation technique proposed as an alternative or complementary treatment for ...

A Preclinical Model to Assess Brain Recovery After Acute Stroke in Rats

The purpose of this study was to establish and validate an animal brain ischemia model in the recovery and sequela stages. A middle cerebral artery occlusion/reperfusion (MCAO/R) model in male Sprague-Dawley rats was chosen. By changing the rat&#39;s weight (260&#8722;330 g), the thread bolt type (2636/2838/3040/3043) and the brain infarct time (2-3 h), a higher Longa&#39;s score, a larger infarct volume and a greater model success ratio were screened using the Longa&#39;s score and TTC staining. The optimum model condition (300 g, 3040 thread bolt, 3 h brain infarct time) was acquired and used in a 1-90 day observation period after reperfusion via assessment of sensorimotor functions and infarct volume. At these conditions, the bilateral asymmetry test had a significant difference from 1 to 90 days, and the grid-walking test had a significant difference from 1 to 60 days; both differences could be a suitable sensorimotor functional test. Thus, the most appropriate condition of a novel rat model in the recovery and sequela stages of brain ischemia was found: 300 g rats that underwent MCAO with a 3040 thread bolt for a 3 h brain infarct and then reperfused. The appropriate sensorimotor functional tests were a bilateral asymmetry test and a grid-walking test.

The&#160;purpose of this study is to establish and validate an animal model for research in the recovery and sequela stages of brain ischemia by testing brain infarction and sensorimotor function after middle cerebral artery occlusion/reperfusion (MCAO/R) after 1-90 days in rats.

The purpose of this study was to establish and validate an animal brain ischemia model in the recovery and sequela stages. A middle cerebral artery ...

Double Direct Injection of Blood into the Cisterna Magna as a Model of Subarachnoid Hemorrhage

Among strokes, subarachnoid hemorrhage (SAH) consecutive to the rupture of a cerebral arterial aneurysm represents 5-9% but is responsible for about 30% of the total stroke-related mortality with an important morbidity in terms of neurological outcome. A delayed cerebral vasospasm (CVS) may occur most often in association with a delayed cerebral ischemia. Different animal models of SAH are now being used including endovascular perforation and direct injection of blood into the cisterna magna or even the prechiasmatic cistern, each exhibiting distinct advantages and disadvantages. In this article, a standardized mouse model of SAH by double direct injection of determined volumes of autologous whole blood into the cisterna magna is presented. Briefly, mice were weighed and then anesthetized by isoflurane inhalation. Then, the animal was placed in a reclining position on a heated blanket maintaining a rectal temperature of 37 &#176;C and positioned in a stereotactic frame with a cervical bend of about 30&#176;. Once in place, the tip of an elongated glass micropipette filled with the homologous arterial blood taken from carotid artery of another mouse of the same age and gender (C57Bl/6J) was positioned at a right angle in contact with the atlanto-occipital membrane by means of a micromanipulator. Then 60 &#181;L of blood was injected in the cisterna magna followed by a 30&#176; downward tilt of the animal for 2 minutes. The second infusion of 30 &#181;L of blood into the cisterna magna was performed 24 h after the first one. The individual follow-up of each animal is carried out daily (careful evaluation of weight and well-being). This procedure allows a predictable and highly reproducible distribution of blood, likely accompanied by intracranial pressure elevation that can be mimicked by an equivalent injection of an artificial cerebral spinal fluid (CSF), and represents an acute to mild-model of SAH inducing low mortality.

We described in this protocol a standardized subarachnoid hemorrhage (SAH) mouse model by a double injection of autologous whole-blood into the cisterna magna. The high degree of standardization of the double-injection procedure represents a middle-to-acute model of SAH with relative safety regarding mortality.

Among strokes, subarachnoid hemorrhage (SAH) consecutive to the rupture of a cerebral arterial aneurysm represents 5-9% but is responsible for about 30% ...

Using a Cell-Tracer Injection to Investigate the Origin of Neointima-Forming Cells in a Rat Saccular Side Wall Model

Microsurgical clipping creates a subsequent barrier of blood flow into intracranial aneurysms, whereas endovascular treatment relies on neointima and thrombus formation. The source of endothelial cells covering the endoluminal layer of the neointima remains unclear. Therefore, the aim of the present study was to investigate the origin of neointima-forming cells after cell-tracer injection in the already well-established Helsinki rat microsurgical sidewall aneurysm model.
Sidewall aneurysms were created by suturing decellularized or vital arterial pouches end-to-side to the aorta in male Lewis rats. Before arteriotomy with aneurysm suture, a cell-tracer injection containing CM-Dil dye was performed into the clamped aorta to label endothelial cells in the adjacent vessel and track their proliferation during follow-up (FU). Treatment followed by coiling (n = 16) or stenting (n = 15). At FU (7 days or 21 days), all rats underwent fluorescence angiography, followed by aneurysm harvesting and macroscopic and histological evaluation with immunohistological cell counts for specific regions of interest. 
None of the 31 aneurysms had ruptured upon follow-up. Four animals died prematurely. Macroscopically residual perfusion was observed in 75.0% coiled and 7.0% of stented rats. The amount of cell-tracer-positive cells was significantly elevated in decellularized stented compared to coiled aneurysms with respect to thrombus on day 7 (p = 0.01) and neointima on day 21 (p = 0.04). No significant differences were found in thrombus or neointima in vital aneurysms. 
These findings confirm worse healing patterns in coiled compared to stented aneurysms. Neointima formation seems particularly dependent on the parent artery in decellularized aneurysms, whereas it is supported by the recruitment from aneurysm wall cells in vital cell-rich walls. In terms of translation, stent treatment might be more appropriate for highly degenerated aneurysms, whereas coiling alone might be adequate for aneurysms with mostly healthy vessel walls.

We performed a one-point, lipophilic cell-tracer injection to track endothelial cells, followed by an arteriotomy and suturing of sidewall aneurysms on the abdominal rat aorta. Neointima formation seemed dependent on the parent artery in decellularized aneurysms and was promoted by the recruitment from aneurysm wall cells in vital cell-rich walls.

Microsurgical clipping creates a subsequent barrier of blood flow into intracranial aneurysms, whereas endovascular treatment relies on neointima and ...