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

Summary

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.

Abstract

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.

Introduction

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's population¹ and causing a substantial socioeconomic burden on society^2,3. Treatment options and their efficacy have been suboptimal and only provide symptomatic relief and do not significantly modify pathophysiological events that underly migraine occurrence^4,5. 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 ganglia^6,7,8,9,10. 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 times^11,12. 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 animals^12,13,14. 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 sufferers^16,17 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 stimuli^18,19. 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 migraine^20,21. 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.

Protocol

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

1. 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).
2. 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.
   1. Take caution not to bend or dull the infuser when adjusting the plastic cap.
   2. 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.
3. 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).
   1. Cut the tubing to a minimum length of 8 in to ensure there is sufficient line to hold a volume of 5 µL.
   2. 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.
4. Attach the other end of the tubing to a 10 µL 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).
5. Once the line is connected, backfill the syringe with 5 µL of phosphate-buffered saline (PBS), synthetic interstitial fluid (SIF), or other vehicles of choice to prevent air bubbles from forming.
   1. 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.
6. After the line is backfilled with 5 µL of the vehicle and is working efficiently, load 5 µL 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).
   1. 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.
7. The maximum volume tested in mice that did not cause leaking into the brain is roughly 10 µL. The behavioral effects after injections with this volume have not been tested. For this reason, administer only 5 µL of the solution onto the dura.
   ​NOTE: These observations are based on the mouse strains/ages/weights of 6-8 week old CD1/ICR mice.

2. Dural injections

1. 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.
   1. After the mouse no longer displays a pinch reflex, adjust the anesthesia and sustain it at a 1.5% isoflurane.
2. 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 in a position conducive to a successful injection.
3. Use one hand to steady the head of the animal and hold the infuser with the other hand.
4. Carefully probe and locate the junction of the sagittal and lambdoidal sutures on the skull of the mouse (Figure 1B, C).
   1. 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.
   2. Verify the position of the junction by re-positioning the infuser along the skull and feeling for the exact location.
5. 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.
6. To verify the accuracy, use ink or dye as an injection solution and euthanize and decapitate the mouse.
   1. 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.
7. 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.
   ​NOTE: Allow the mouse to recover from anesthesia for a minimum of 30 min prior to doing any behavioral experiments.

3. Periorbital von Frey

1. Begin the study with a cohort of approximately 16-20 mice.
2. One the day prior to habituation, handle each mouse for at least 5 min.
3. 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.
   1. 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).
4. 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.
   1. Ensure that every time the mice are in the chamber that there is access to food.
   2. 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.
5. Following the initial 3 days of habituation, place the mice in their individual chambers.
   1. 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.
6. After acclimation to the room on the testing day, remove one mouse while still in its cup from its respective chamber.
   1. 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's responses and even prevent animals from responding.
7. Place the cup with the mouse inside on the table below the testing rack on the absorbent pad.
8. For periorbital von Frey testing, place the 0.07 g von Frey filament directly in the center of the face and between the eyes.
9. Apply enough pressure on the filament to cause the von Frey hair to bend into a "C" shaped formation.
   1. 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.
   2. Apply von Frey filaments according to the Dixon "up-down" method^22,23.
      1. 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.
      2. 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.
      3. In this method, if an animal does not exhibit a response to the filament, apply the filament of the next higher gram weight.
      4. 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.
      5. 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.
         ​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.

4. Testing for baseline withdrawal thresholds

1. Prior to inclusion in an experiment, ensure that the mice reach a baseline withdrawal threshold between 0.5-0.6 g.
   1. 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).
2. Test mice daily when establishing baseline withdrawal thresholds.
   1. Testing allows animals to acclimate to the testing conditions and the pressure of the von Frey filaments.
   2. 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.
3. Test mice for approximately 7 days before determining which animals do not meet the inclusion criteria for an experiment.
   ​NOTE: Approximately 70% of mice will reach the targeted baseline level.
   1. 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.
   2. 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.

5. Analysis of von Frey results

1. 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.
2. 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.
3. Plot the data as either 50% withdrawal threshold or average withdrawal threshold in grams.

Results

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 Frey^12,13,14. Here we show how this model can be used to assess potential sex-specific contributions to migraine pathology (Figure 3).

Thi...

Discussion

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 injury^25,26. 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...

Materials

List of materials used in this article

Name	Company	Catalog Number	Comments
4 oz Hot Paper Cups	Choice Paper Company	5004W	https://www.webstaurantstore.com/choice-4-oz-white-poly-paper-hot-cup-case/5004W.html
Absorbent Underpads	Fisherbrand	14-206-65	https://www.fishersci.com/shop/products/fisherbrand-absorbent-underpads-8/p-306048
C313I/SPC Internal 28 G cannula	P1 Technologies (formerly Plastics One)	8IC313ISPCXC	I.D. 18 mm, O.D. 35 mm
Gastight Model 1701 SN Syringes	Hamilton	80008	https://www.hamiltoncompany.com/laboratory-products/syringes/80008
Ismatec Pump Tubing, 0.19 mm	Cole-Palmer	EW-96460-10	https://www.coleparmer.com/i/ismatec-pump-tubing-2-stop-tygon-s3-e-lab-0-19-mm-id-12-pk/9646010
Stand with chicken wire	Custom		The galvanized steel chicken wire dimensions are 0.25 in. x 19-gauge
Testing Rack with individual  Chambers	Custom		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.
von Frey Filaments	Touch test/Stoelting	58011	https://www.stoeltingco.com/touch-test.html