Modeling Multiple Sclerosis in the Two Sexes: MOG35-55-Induced Experimental Autoimmune Encephalomyelitis

Summary

Experimental autoimmune encephalomyelitis is one of the most widely used murine models of multiple sclerosis. In the current protocol, C57BL/6J mice of both sexes are immunized with myelin oligodendrocyte glycoprotein peptide, resulting mainly in ascending paresis of the tail and limbs. Here we discuss the protocol of EAE induction and evaluation.

Abstract

Multiple Sclerosis (MS) is a chronic autoimmune inflammatory disease affecting the central nervous system (CNS). It is characterized by different prevalence in the sexes, affecting more women than men, and different outcomes, showing more aggressive forms in men than in women. Furthermore, MS is highly heterogeneous in terms of clinical aspects, radiological, and pathological features. Thus, it is necessary to take advantage of experimental animal models that allow the investigation of as many aspects of the pathology as possible. Experimental autoimmune encephalomyelitis (EAE) represents one of the most used models of MS in mice, modeling different disease features, from the activation of the immune system to CNS damage. Here we describe a protocol for the induction of EAE in both male and female C57BL/6J mice using myelin oligodendrocyte glycoprotein peptide 35-55 (MOG_35-55) immunization, which leads to the development of a chronic form of the disease. We also report the evaluation of the daily clinical score and motor performance of these mice for 28 days post immunization (28 dpi). Lastly, we illustrate some basic histological analysis at the CNS level, focusing on the spinal cord as the primary site of disease-induced damage.

Introduction

Multiple Sclerosis (MS) is a chronic autoimmune inflammatory disease affecting the central nervous system (CNS). It shows the presence of perivascular infiltration of inflammatory cells, demyelination, axonal loss, and gliosis¹. Its etiology remains unknown, and its clinical aspects, radiographic, and pathological features suggest remarkable heterogeneity in the disease².

Due to its unknown etiology and complexity, at present, no animal model recapitulates all the clinical and radiological features displayed in human MS^3,4. However, various animal models are employed to study different aspects of MS^3,4. In these models, disease initiation is typically extremely artificial, and the timeframe of the onset of clinical signs is different between humans and mice. For example, in humans, the pathophysiological processes underlying the disease are undetected for years before the onset of clinical manifestations. Conversely, the experimenters can detect symptoms in animal models within weeks or even days after MS induction⁴.

Three basic animal models produce the features of demyelination that are characteristic of MS: those that are virus-induced (e.g., Theiler's murine encephalomyelitis virus), those that are induced by toxic agents (e.g., cuprizone, lysolecithin), and the different variants of experimental autoimmune encephalomyelitis (EAE)⁵. Each model helps study some specific facets of the disease but none replicates all the features of MS⁶. Thus, it is critical to choose the correct model considering the specific experimental needs and the scientific questions to be addressed.

Thanks to immunization procedures against myelin-derived antigens, EAE is induced by triggering an autoimmune response to CNS components in susceptible mice. The interplay between a wide range of immunopathological and neuropathological mechanisms causes the development of principal pathological traits of MS (i.e., inflammation, demyelination, axonal loss, and gliosis) in the immunized mice^7,8. Mice begin to show clinical symptoms around the second week after immunization and generally show ascending paralysis from the tail to the limb and forelimb. The clinical score (i.e., quantification of the accumulation of disease-related deficits) is generally assessed using a 5-point scale⁷.

Active immunization with protein or peptide or passive transfer of encephalitogenic T cells can be used to induce EAE in mice with different genetic backgrounds (e.g., SJL/J, C57BL/6, and non-obese-diabetic (NOD) mice). Myelin proteolipid protein (PLP), myelin basic protein (MBP), and myelin oligodendrocyte glycoprotein (MOG) are examples of self-CNS proteins from which immunogens are usually produced. Particularly, SJL/J mice immunized with the immunodominant epitope of PLP (PLP_139−151) develop a relapsing-remitting (RR) disease course, while C57BL/6J mice immunized with the immunodominant MOG_35-55 peptide show EAE of a chronic nature¹. Despite some limitations, such as providing very little information about MS progression, the role of B cells in the disease, the inside-out mechanisms, or difficulties in studying remyelination, the EAE models have hugely contributed to the understanding of autoimmune and neuroinflammatory processes, increasing the knowledge in the MS field and thus allowing the development of novel therapeutic approaches for this disease^4,6.

In the present work, we focused on a particular form of active EAE, the myelin oligodendrocyte glycoprotein peptide 35-55 (MOG_35-55)-induced form^9,10,11,12. The MOG_35-55-induced EAE models a chronic form of MS. After immunization, the mice undergo an asymptomatic phase within the first week after immunization, then the disease typically arises during the second week after immunization, while between the third and fourth weeks after immunization, the disease becomes chronic, with no possibility of full recovery from the accumulated deficits^7,8,13. Interestingly, no differences between males and females in incidence, disease onset, course, or progression are observed in most of the studies present in the literature¹⁴, even if fewer studies compare the disease in males and females.

In contrast, in humans, these parameters are known to be strongly sexually dimorphic². MS affects more women than men; however, men generally develop a more aggressive form of the disease². This evidence has suggested an essential, as well as complex, role of the gonadal hormones¹⁵; nevertheless, the role and the mechanism of action of sex hormones in the pathology remain unclear. Moreover, data from animal models support the idea that both estrogens and androgens exert positive effects on different tracts of the pathology in a sex-specific manner^16,17.

Some studies also suggest neuroprotective, promyelinating, and anti-inflammatory effects of progesterone¹⁸ and, although evidence in MS patients is scarce¹⁸, neuroactive steroids (i.e., de novo synthetized steroids by the nervous system, such as pregnenolone, tetrahydroprogesterone, and dihydroprogesterone) might also affect the pathological course¹⁹. Collectively, these data support the idea that sex hormones produced both peripherally and inside the CNS have an important and sex-specific role in disease onset and progression. Therefore, in the present work, we urge the collection of separate data from both male and female animals.

From the histopathological point of view, the white matter of the spinal cord serves as the principal site of CNS injury in this model, which is characterized by multifocal, confluent regions of mononuclear inflammatory infiltration and demyelination⁸. Thus, in describing this protocol for the induction of MOG_35-55-induced EAE in C57BL/6J mice, we will take into account the disease outcome in the two sexes and provide some histopathological insights regarding the spinal cords.

Protocol

The animal care and handling in the present work was performed according to the European Union Council Directive of 22^nd September 2010 (2010/63/UE); all the procedures reported in the present study were approved by the Italian Ministry of Health (407/2018-PR) and by the Ethical Committee of the University of Torino (Project n° 360384). We suggest conforming to the experimental design to the ARRIVE guidelines originally published by Kilkenny et al. in 2010²⁰. Before starting, ensure that the necessary materials are available (see Table of Materials). Sterilize all glassware and utensils used for the preparation of the MOG_35-55 emulsion in an autoclave. A summary of the experimental procedures is represented in Figure 1.

1. Preparation of MOG_35-55 emulsion

NOTE: To prepare the emulsion, MOG_35-55, incomplete Freud's adjuvant (IFA), Mycobacterium Tuberculosis strain H37Ra (MT), and physiological solution are required (see Table of Materials).

CAUTION: Heat-killed MT can stimulate the innate immune response. Avoid inhalation, ingestion, and contact with skin and eyes using proper personal protective equipment and weighing MT in a covered precision balance under the hood.

Solution	Composition	Notes
2 mg/mL MOG35-55 peptide  solution	Lyophilized MOG35-55 peptide diluted in physiological solution at 2 mg/mL concentration	Preserve the already diluted solution at -80 °C.
5 µg/mL PT solution	Lyophilized PT diluted in physiological solution at 5 µg/mL concentration.	Preserve the already diluted solution at -80 °C.
Emulsion	The total volume of emulsion needed for each mouse to be immunized is 300 µL divided as follows:	To avoid alteraions or contamination, prepare the emulsion the day of the immunization.
	200 µg/mouse of MOG35-55 , i.e., 100 µL of MOG35-55  2 mg/mL solution.
	50 µL of physiological solution
	150 µL of IFA
	4 mg/mL MT, i.e., 1.2 mg/mouse
Physiological solution	Sodium chloride 0.9% diluted in distilled water.

Table 1: Composition of the solutions used for the immunization procedure.

1. Prepare the solution in a glass beaker, adding the liquid component first and finally the MT.
   NOTE: The total volume of emulsion needed for each mouse to be immunized is 300 µL, divided as indicated in Table 1. The emulsion is very viscous and thick, so during the preparation and injection procedure, there could be some loss, especially when preparing the solution for a few mice. We suggest calculating the final volume of emulsion needed by overestimating the number of mice to be immunized by at least 1.5-2-fold.
2. Place the beaker in ice and use the glass syringe with an 18 G needle to start emulsifying the solution.
   NOTE: The emulsion can also be prepared using other strategies, for example, by connecting the two air-free glass syringes with a three-way stopcock and mixing the solution by pushing the plungers back and forth^21,22.
3. Emulsify the solution for at least 15 min; generally, 30 min of emulsification is enough. To check the quality of the emulsion, add a drop of emulsion in a transparent container filled with water: if the drop maintains its structure and remains intact, the emulsion is ready.
4. Place the emulsion directly inside the 1 mL syringes that will be used for the immunization and store these at +4 °C until use to preserve the thickness of the emulsion and avoid alterations or contaminations.

2. Animal selection and immunization

1. Animal selection
   1. Select adult C57BL/6J mice of both sexes at 8-10 weeks of age with an optimal body weight of ~20 g. Be sure to select age- and sex-matched mice for different experimental groups because the susceptibility to disease can vary with age and sex.
      NOTE: Mice should be of comparable body weight on the day of immunization because the present procedure is optimized for a certain body weight range (17-25 g).
   2. House same-sex animals in groups (n = 4-5/cage) to avoid social isolation in standard conditions in 45 cm x 25 cm x 15 cm polypropylene mouse cages at 22 ± 2 °C, under 12:12 light/dark cycle (lights on at 08:00 AM). Provide food and water ad libitum.
      NOTE: Eventually, to further limit the potential variability of disease course in the immunized animals, we also suggest forming sufficiently numerous experimental groups. There are also studies^8,23 that describe the timing of immunization as a condition that determines variation in EAE outcome. Thus, we suggest performing the immunization approximately at the same hour in all animals, preferably during the light hours of the daily light/dark cycle²³. To limit stress, it is preferable to manipulate the animals prior to the day of immunization and to mark them to easily identify them for daily evaluation, for example, ear clipping or tag.
2. The immunization procedure
   NOTE: Ensure that the procedure is performed by an experienced investigator to minimize stress for animals and optimize immunization. Before starting the immunization, select the anesthesia method in accordance with the institutional animal care and ethical committee guidelines. Our laboratory uses brief anesthesia with isoflurane.
   1. Anesthetize the mouse: 4% isoflurane for anesthesia induction and 1.5-2% isoflurane for anesthesia maintenance. Wait for the anesthesia to be effective and use a back and front foot toe pinch to assess the level of anesthesia. To prevent dryness of the eyes while the mouse is under anesthesia, use a vet-approved eye ointment.
   2. PT intravenous injection in the lateral caudal vein: plug the tail of the mouse gently with an ethanol solution, which acts as a vasodilator, to display the veins. Focus on one of the two lateral veins of the tail, and, using a 0.5 mL syringe fitted with a 30 G needle, inject 500 ng of PT (i.e., 100 µL of PT diluted in physiological solution at the concentration of 5 µg/mL).
   3. Subcutaneous injection of MOG_35-55 emulsion: using the 1 mL syringe with a 26 G needle, perform three subcutaneous injections of the previously prepared emulsion: two under the rostral part of the flanks and one at the base of the tail. The volume of emulsion injected in each site is 100 µL, for a total volume of 300 µL of emulsion injected in each mouse.
      ​NOTE: The day of immunization is recorded as day 0 post immunization (dpi); 48 h later (i.e., at 2 dpi), it is necessary to perform another intravenous injection of PT equal to the one previously performed. It is possible to perform the injection without anesthesia, using a mouse restrainer. The procedure described here aims to induce and maintain anesthetizing of the mice during the immunization procedure to avoid possible discomfort and movements of the animals or risks for both mice and investigators. Thus, there are no surgical procedures. However, to optimize the experimental processes, it is important to maintain appropriate sterile conditions during these steps and to monitor the mouse's condition after these procedures.
   4. To verify the mouse has recovered from the injections, place it in a clean cage after the procedures and wait until it has regained sufficient consciousness to maintain sternal recumbency. After the mouse has fully recovered, return it to the home cage with other animals.

3. EAE follow-up

1. Body weight and food intake
   1. Monitor daily the body weight (BW) of the animals, using an electronic precision balance, because the decrease in BW is an indicator of disease progression.
      NOTE: This decrease should not exceed a certain percentage, according to the institutional and ethical committee's guidelines for animal care. Usually, if an animal loses more than 20% of the initial body weight (i.e., the weight recorded at 0 dpi), it should be sacrificed as the application of the humane endpoint. The euthanasia methods involve deep irreversible anesthesia for inhalation (e.g., 5% isoflurane) followed by decapitation.
   2. Monitor the food intake (FI)-the food eaten by an animal in a day (g∙day^-1∙animal^-1), weighing the amount of food in the specific container at least once a week, and dividing the amount of eaten food for the days passed between two sequential measurements and the number of animals present in the cage.
      ​NOTE: This measurement allows the estimation of mean food intake. Because of the appearance and accumulation of clinical disease signs, which involve the paralysis of the limbs, we suggest placing some watered food on the floor of the cage when the animals are not capable of standing firmly on their hind limbs and reaching the food container or the water bottle. To assess the food intake through the follow-up period as precisely as possible, we also measured the dry weight of this food before wetting it for the mice.
2. Evaluation of estrous cyclicity during EAE
   1. Check the estrous cycle for at least two cycles, evaluating the vaginal cytology smears as described by McLean et al.²⁴. Classify the phase of the estrous cycle based on the presence of three primary cell types-nucleated epithelial cells, cornified squamous epithelial cells, and leukocytes-in the vaginal smear samples, as follows:
      1. Classify as proestrus based on an almost exclusive presence of clusters of round, well-formed nucleated epithelial cells.
      2. Classify as estrus based on the predominant presence of densely packed clusters of cornified squamous epithelial cells.
      3. Classify as metestrus based on the predominant presence of small darkly stained leukocytes and the minor presence of cornified squamous epithelial cells.
      4. Classify as diestrus based on the highly predominant presence of small darkly stained leukocytes and rare cornified squamous epithelial cells along with the possible appearance of nucleated epithelial cells.
         NOTE: When evaluating the disease in both sexes, it is important to check the variability in females due to the estrous cycle. We suggest focusing on the evaluation of the estrous cycle, especially between the first and the second-week post immunization (i.e., during the acute phase of the EAE). It has been previously shown that the immunization procedure causes the most pronounced alterations of the estrous cycle within this phase²⁵. Moreover, it is important to consider that performing the smear when the animal reaches a high clinical score (especially >3) is difficult due to the posterior paresis and the lack of tone in the hindlimbs.
3. Clinical score
   1. Have a blinded investigator assess the clinical score of the animals daily. Assign to each animal a score rated from 0 to 5 (see Table 2) to evaluate the disease course²⁶ as described by Racke⁷.
      NOTE: Similar to the body weight decrease, a humane endpoint is also necessary for the increase in clinical scores, according to the institutional and ethical committee's guidelines for animal care. Generally, if an animal is no longer able to feed itself autonomously (this usually occurs when an animal reaches at least the score of 4, according to the scale we use), it should be sacrificed as the application of the humane endpoint.
4. Motor performance evaluation by rotarod test
   NOTE: Evaluation of the EAE progression is usually performed by assigning the clinical score daily, which is done by a fully trained blinded investigator. However, it could be useful to flank it with a more quantitative and objective evaluation of the disease's progression. In a previous study²⁶, the rotarod test was used to measure the motor performance of the immunized animals. As described by van den Berg et al.²⁷, to have a more quantitative and precise clinical evaluation of the disease course, the evaluation of the motor performance by the rotarod test can support the clinical score assessment. For a detailed description, see van den Berg et al.²⁷.
   1. Let the mice undergo rotarod sessions daily, starting from 1 dpi until the sacrifice (i.e., 28 dpi). Each session consists of a single 300 s session during which the rod speed has to be increased linearly from 4 to 40 rpm.
   2. Register the animal's score. When the mouse is not capable of maintaining its balance and falls off the device, it falls on the ground and triggers a sensor, and the time (s) is recorded. Thus, the performance is scored as latency to fall (s).

Grade	Clinical sign		Description
0	Healthy		No observed clinical sign. The animal shows a normal tone and moving of the tail. It walks without tripping.
0.5	Impaired gate		The animal trips while walking on a grill.
1	Limp tail		When the animal is picked up by the basis of the tail, the tail droops (flabby tail).
1.5	Limp tail and impaired gate		The animal shows a flabby tail, and it trips while walking on a grill.
2	Ataxia		The animal displays difficulties in standing up once it has been turned on its back.
2.5	Ataxia and paresis of hindlimb		The animal displays cannot stand up once it has been turned on its back, and it loses the tone of one of its hindlimbs.
3	Paralysis of hindlimbs		The animal loses the tone of both hindlimbs.
3.5	Paralysis of hindlimbs and/or paresis of forelimb		The animal loses the tone of both hindlimbs and partially of forelimbs. In fact, it shows a loss of strength in the forelimbs’ grasp.
4	Tetra paresis		The animal completely loses the tone of its limbs.
4.5	Tetra paresis and decreased body temperature		The animal completely loses the tone of its limbs, and it shows a decrease in body temperature (it is cold).
5	Dying or dead		The animal is dying (it does not respond to any stimulus) or dead.

Table 2: Clinical scoring system used to assess EAE progression.

4. Evaluation of EAE-induced histopathological signs at the spinal cord level

NOTE: Here, we briefly report the procedure to sacrifice the animals and collect the spinal cords to perform histopathological analysis; for a detailed description, see these references^10,26,28,29.

1. Fixation and tissue sampling
   NOTE: For a detailed description, see these references^10,26.
   1. Sacrifice the animals at 28 dpi during the chronic phase of the disease.
      1. Anesthetize the mice by deep irreversible anesthesia (intraperitoneal injection of Zolazepam and Tiletamine 80 mg/kg / Xylazine 10 mg/kg).
      2. Transcardially perfuse the mice with a saline solution followed by a 4% paraformaldehyde (PFA) solution.
         NOTE: Pay attention while using PFA: as it is toxic, avoid inhalation, ingestion, and contact with skin and eyes using proper personal protective equipment. Weigh the powder, prepare the solution, and perform the perfusion under the hood.
   2. Remove the spinal cords from the spinal column³⁰.
   3. Store the spinal cords in a 4% PFA solution for 24 h.
   4. Perform several washes in 0.01 M saline phosphate buffer (PBS).
   5. Embed the spinal cords in paraffin blocks³⁰.
2. Histological procedures
   NOTE: For a detailed description, see Montarolo et al.¹⁰.
   1. Use a microtome to cut 10 µm thick transverse spinal cord sections and collect them on gelatin-coated slides. Orient the plane of sectioning to match the drawings corresponding to the transverse sections of the mouse spinal cord atlas³¹.
   2. Perform the deparaffinization of the sections³².
   3. Stain the section with Hematoxylin and Eosin³².
   4. Dehydrate the sections³².
   5. Cover the sections with a mounting medium and let them dry at room temperature under a chemical hood.
      NOTE: Hematoxylin-Eosin staining allows for the detection of the presence of the perivascular inflammatory infiltrates (PvIIs)²⁶, which is assessed as a sign of the disease²⁸.
3. Quantitative analysis of spinal cord sections
   1. Acquire images of the stained sections with an optical microscope connected to a digital camera with a 20x objective²⁹.
   2. Analyze the acquired image to obtain the number of PvIIs, expressed as the number of infiltrates per mm².
      NOTE: For the analysis of the acquired images, it is useful to take advantage of image analysis software. Neuropathological findings presented in this work are quantified in 10 complete cross-sections of the spinal cord per mouse (n = 8/group) representative of whole spinal cord levels.

Results

EAE follow-up after immunization
This was assessed as described below.

Body weight and food intake
The two-way analysis of variance (ANOVA) (sex and time as independent variables) shows a decrease in the BW of EAE animals of both sexes, especially within the second week post induction (F_(1,57) = 4.952, p < 0.001; Figure 2A). However, the sexual dimorphism in BW is always maintained (

Discussion

The MOG_35-55-induced EAE protocol that we described led to the development of a chronic form of MS in C57BL/6J mice^7,8,13. In these representative results, we reported that the animals of both sexes that underwent the immunization procedure developed a chronic form of the disease (i.e., they do not fully recover after the disease onset, they accumulate deficits, and maintain a CS at least of 1.5 in the chronic phase)....

Materials

Name	Company	Catalog Number	Comments
18 G x 1 ½“ 1.2 x 40 mm needle for the glass syringe	Terumo	TER-HYP-18G-112-PIN
Digital camera connected to the optical microscope	NIKON DS-U1 digital camera
Electronic precision balance	Merck	Mod. Kern-440-47N, resolution 0.1 g
Eosin Y	Sigma-Aldrich	HT110216
Glass syringe pipet “ultra asept” 10 ml	Sacco System	L003465
Glassware (i.e., becker to prepare the emulsion)	VWR	213-1170, 213-1172
Hematoxylin (Mayer’s)	Sigma-Aldrich	MHS32	Filter before using it.
Image analysis Software	Fiji
Incomplete Freund’s adjuvant (IFA)	Sigma-Aldrich	F5506	Store at +4 °C.
Isoflurane	Wellona Pharma		This drug is used as inhalational anaesthetic.
Male and female C57BL/6J mice	Jackson Laboratory, Envigo		Age 8-10 weeks, optimal body weight of ~20 g.
Microtome	Leica HistoCore BIOCUT R
Mounting Medium	Merck	107961
Mouse Rotarod	Ugo Basile	#47600
Mycobacterium tuberculosis (MT), strain H37Ra	Difco Laboratories Inc.	231141	Store at +4 °C.
Myelin oligodendrocyte glycoprotein peptide 35-55 (MOG35-55)	Espikem	EPK1	Store at -80 °C diluted (2 mg/mL) in physiological solution; prepare it on the day of the immunization to avoid, as much as possible, alterations or contaminations.
Optical microscope	NIKON eclipse 90i
Paraformaldehyde (PFA)	Sigma-Aldrich	158127	Store at +4 °C once diluted (4%) in phosphate buffer.
Pertussis toxin (PT)	Duotech	PT.181	Store at -80°C diluted (concentration 5 µg/mL) in physiological solution
Physiological solution (sodium chloride 0.9% solution)	B. Eurospital	A 032182038	Store at +4 °C once opened.
Saline phosphate buffer (PBS)	Thermo Scientific	J61196.AP
Software for image acquisition	NIS-Element AR 2.10
Syringes U-100 0.5 mL with 30 G x 5/16” (0.30 x 8 mm) in fixed needle	Nipro	SYMS-0.5U100-3008B-EC
Syringes U-100 1 mL with 26G x ½” (0.45 x 12.7 mm) in needle	PIC	20,71,26,03,00,354
Vet ointment for eyes	Lacrilube, Lacrigel Europhta
Xylazine	Rompun		This mixture of drug is used as injectable anaesthetic and sedative.
Zolazepam and Tiletamine	Zoletil  100		This drug is used as injectable anaesthetic, sedative, muscle relaxer, and analgesic