Transcranial Direct Current Stimulation (tDCS) in Mice

Podsumowanie

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.

Streszczenie

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.

Wprowadzenie

Transcranial Direct Current Stimulation (tDCS) is a non-invasive, low-cost, therapeutic technique, which focuses on neuronal modulation through the use of low-intensity continuous currents¹. There are currently two setups (anodal and cathodal) for tDCS. While the anodal stimulation exerts a current electric field too weak to trigger action potentials, electrophysiology studies have shown that this method produces changes in synaptic plasticity². For example, evidence shows that tDCS induces long-term potentiation (LTP) effects such as increased peak amplitude of the excitatory postsynaptic potentials^3,4 and modulation of cortical excitability⁵.

Conversely, cathodal stimulation induces inhibition, resulting in membrane hyperpolarization⁶. A hypothesis for this mechanism is based on the physiological findings where tDCS is described to modulate action potential frequency and duration in the neuronal body³. Notably, this effect does not directly evoke action potentials, though it can shift the depolarization threshold and facilitate or hamper neuronal firing⁷. These contrasting effects have been previously demonstrated. For example, anodal and cathodal stimulation produced opposing effects in conditioned responses registered via electromyography activity in rabbits⁸. However, studies have also shown that prolonged anodal stimulation sessions may decrease excitability while increasing cathodal currents may lead to excitability, presenting self-contrasting effects³.

Both anodal and cathodal stimuli aggregate the use of electrode pairs. For example, in anodal stimulation, the "active" or "anode" electrode is placed over the brain region to be modulated whereas the "reference" or "cathode" electrode is situated over a region where the effect of current is assumed to be insignificant⁹. In the cathodal stimulation, electrode disposition is inverted. The stimulation intensity for effective tDCS depends on the current intensity and electrode dimensions, which affect the electric field differently¹⁰. In most published studies, the average current intensity is between 0.10 to 2.0 mA and 0.1 mA to 0.8 mA for human and mice, respectively^6,11. Although the electrode size of 35 cm² is typically used in humans, there is no proper understanding regarding electrode dimensions for rodents and a more thorough investigation is needed⁶.

tDCS has been proposed in clinical studies with the attempt of offering an alternative or complementary treatment for several neurological and neuropsychiatric disorders¹¹ such as epilepsy¹², bipolar disorder¹³, stroke⁵, major depression¹⁴, Alzheimer's disease¹⁵, multiple sclerosis¹⁶ and Parkinson's disease¹⁷. Despite growing interest in tDCS and its use in clinical trials, detailed cellular and molecular evoked alterations in brain tissue, short and long-lasting effects, as well as behavioral outcomes, are yet to be more deeply investigated^18,19. Since a direct human approach to thoroughly study tDCS is not viable, the use of a tDCS animal model may offer valuable insights into the cellular and molecular events underlying the therapeutic mechanisms of tDCS due to the accessibility to the animal's brain tissue.

Available evidence is limited regarding tDCS models in mice. Most of the reported models used different implanting layouts, electrode dimensions, and materials. For example, Winkler et al. (2017) implanted the head electrode (Ag/AgCl, 4 mm in diameter) filled with saline and fixed it to the cranium with acrylic cement and screws²⁰. Different from our approach, their chest electrode was implanted (platinum, 20 x 1.5 mm). Nasehi et al. (2017) used a procedure very similar to ours, although the thoracic electrode was made from a saline-soaked sponge (carbon filled, 9.5 cm²)²¹. Another study implanted both electrodes into the animal's head, which was achieved by using fixed plates and covering the animal's head with a hydrogel conductor²². Here, we describe a tDCS mouse model that uses a chronically implanted electrode through simple surgical procedures and tDCS setup (Figure 1).

Protokół

Individually-housed male adult (8-12 weeks) C57BL/6 mice were used in this experiment. Animals received proper care before, during and after experimental procedures with food and water ad libitum. All procedures were approved by the animal ethics committee from Federal University of Minas Gerais (protocol number 59/2014).

1. Electrode Placement

1. Sedating and fixating the animal onto the stereotaxic apparatus
   1. Sterilize all the necessary surgical instruments.
      Note. Surgical instruments were sterilized for 3 minutes at 440 °C. Cotton swabs were autoclaved at 20 psi (pounds per square inch) at 121 °C for 20 min.
   2. Adjust the thermal platform controller to 37 °C.
   3. Weigh the animal and calculate the appropriate dose for anesthesia induction. Use a mixture of ketamine and xylazine at a dose of 100 mg/kg ketamine and 8 mg/kg xylazine, given intraperitoneally (needle size, 31 G). The animal should fall asleep within 2 to 3 min.
   4. Use an electric shaver or razor to shave down the surgical site.
   5. Place the animal onto the stereotaxic apparatus over the pre-warmed heating plate.
   6. Hold the animal's head and insert the tip ear bars into each of the animal's ears to fix it to the stereotaxic platform.
   7. Verify there is no lateral head shifting and little vertical movement after by slowly shifting the animal's head positioning.
   8. Gently slide the anesthesia mask over the mouse's nose and fix it in place by tightening the screw.
   9. Set the isoflurane to 1% with 1.0 L/min of O₂.
   10. Apply eye ointment to the animal's eyes to prevent corneal drying during the surgery.
2. Attaching the implant to the animal's head
   1. Use the cotton swabs to prepare the surgical site with three alternating scrubs of povidone-iodine (or 2% chlorhexidine) and 70% ethanol.
   2. Use a pair of tweezers to verify anesthesia depth by lightly squeezing the animal's toes and verifying the loss of animal's pedal withdrawal (toe pinch) reflex.
   3. Make an incision about 3 mm posterior to the animal's ear line and stop at the eye line. The incision site must have approximately 1 cm in length to be large enough to receive the implant.
   4. Gently scrape the cranium with a bone scraper to improve glue and cement adherence. Do this light handedly with the intention of creating micro scratches.
   5. Carefully position surgical hooks to the loose skin to maintain an open surgical field and free of obstructions such as skin and fur.
   6. Use a sterile cotton swab to dry the animal's scalp.
   7. Use a dissecting microscope to visualize the top of the animal's cranium.
   8. Attach a needle to the stereotaxic holder and locate the bregma. Position the needle directly above the animal's head slightly touching the bregma.
   9. Zero out all coordinates on the digital tracer and then raise the needle.
   10. Fix the tDCS implant on the stereotaxic holder. Position the implant over the animal's head and lower it slowly onto the region of interest using the proper stereotaxic coordinates.
   11. Use a needle to spread 1 drop (approximately 35 μL) of super glue onto the implant's base.
   12. Slowly move the holder downwards until it touches the skull. Be sure that the implant base is entirely in contact with the surface.
   13. Prepare the surgical cement according to the manufacturer's instructions.
   14. After precise positioning, apply 3 thin, even layer of cement across the cranium and onto the lower portion of the implant. Apply drop per drop using an application brush. Layers must form a hill-shaped structure for further structural support of the implant.
   15. Leave the implant's screw thread clean of cement to allow a smooth, unobstructed connection.
   16. Allow each layer to dry for approximately 4 minutes.
   17. When dry, carefully remove the holder until it is completely detached from the implant. Always use extreme caution when handling the implant, since it may be accidentally extracted from the animal's skull.
3. Finishing surgery and post-surgical care
   1. Hydrate the animal's skin in the incision site with a saline-soaked cotton swab.
   2. Coat the skin over the base of the tDCS implant.
   3. Use a pair of tweezers to bring the tissue together and close the incision with a drop of surgical tissue glue per 0.2 cm of tissue.
   4. Infiltrate 1-2% lidocaine in the incision site and underlying tissues.
   5. Hydrate the mouse with 500 µL of lactate Ringer's solution subcutaneously.
   6. Place the mouse into a pre-warmed (37 °C) clean, single-housed cage.
   7. Put a small dish with wet food pellets in the cage for easy access to food in the following hours.
   8. Register the animal's post-surgical weight.
   9. Give the animal ketoprofen (5 mg/kg) subcutaneously after the surgery and on the next 2 days.
   10. Monitor the recovery of the animal closely for at least 1 week. Assess any sign of distress, such as piloerection, lack of grooming, reduced locomotion, wound scratching and inflammation of the surgical site.

2. tDCS Setup and Stimulation

1. tDCS Setup (see Figure 2)
   ​Note. Make sure that the tDCS stimulator is fully charged.
   1. Attach the anode and cathode cables to the tDCS stimulator and make them available near the stimulation site. Attach the pin-type electrode to the stereotaxic holder.
   2. Set the thermal platform to 37 °C.
   3. Turn on the oxygen flowmeter on the inhalation anesthesia system to 1 L/min.
   4. Place the mouse into the anesthesia induction chamber.
   5. Turn on the isoflurane vaporizer to 3%. Allow the animal to undergo isoflurane effects for 4 min.
   6. While the animal is in the induction chamber, use a sterile syringe to fill the body electrode with 0.9% saline solution.
   7. Remove the animal from the induction chamber and position its chest over the body electrode.
   8. Gently slide the anesthesia mask over the mouse's nose and fix it in place. Lower the isoflurane output to 1.5%.
   9. Fill the implant and the pin-type electrode with saline and carefully attach them.
   10. Adjust stimulation time and current intensity.
   11. Verify the contact quality on the tDCS stimulator.Optimal contact goes from 7 to 10 on a 1 to 10 scale.
2. Stimulation
   1. Start the stimulation.
   2. Observe the current ramping up for 30 s to the selected value and maintaining itself steady for the established time, then, at the end of the session ramping down again.
   3. Activate the sham button for control mice.
   4. Observe the current ramping up for 30 s to the selected value and then down to 1 for the rest of the stimulation period with a final ramp to the selected value at the end with a consecutive ramp down.
   5. Once the stimulation session is complete, carefully transfer the animal to a pre-warmed (37 °C) cage for 10 min.
      Note. Animals start to awaken after 3 min.
   6. Turn the inhalation anesthesia system off.

Wyniki

The surgical protocol presented long-term implant stability for at least one month, with no inflammatory signals at the stimulated site nor any other undesired effect. All the animals survived the surgical procedure and tDCS sessions (n = 8). In this experiment, tDCS implants were positioned over the M1 and M2 cortices (+1.0 mm anterior-posterior and 0.0 mm lateral to bregma). One week later, tDCS (n = 3-4) and sham (n = 3) mice were stimulated for five consecutive days during 10 min at 0...

Dyskusje

In recent years, neurostimulation techniques have been entering clinical practice as a promising procedure to treat neuropsychiatric disorders²³. To reduce the constraint imposed by the lack of knowledge of the mechanisms of neurostimulation, we presented here a tDCS mouse model carrying an electrode that can target brain regions. Since the electrode is chronically implantable, this animal model enables the investigation of long-lasting biological effects evoked by tDCS (for at least 1 month) in c...

Materiały

Name	Company	Catalog Number	Comments
BD Ultra-Fine 50U Syringe	BD	10033430026	For intraperitonially injection.
Shaver (Philips Multigroom)	Philips (Brazil)	QG3340/16	For surgical site trimming.
Surgical Equipment
Model 940 Small Animal Stereotaxic Instrument with Digital Display Console	KOPF	940	For animal surgical restriction and positioning.
Model 922 Non-Rupture 60 Degree Tip Ear Bars	KOPF	922	For animal surgical restriction and positioning.
Cannula Holder	KOPF	1766-AP	For implant positioning.
Precision Stereo Zoom Binocular Microscope (III) on Boom Stand	WPI	PZMIII-BS	For bregma localization and implant positioning.
Temperature Control System Model	KOPF	TCAT-2LV	For animal thermal control.
Cold Light Source	WPI	WA-12633	For focal brightness
Tabletop Laboratory Animal Anesthesia System with Scavenging	VetEquip	901820	For isoflurane delivery and safety.
VaporGuard Activated Charcoal Adsorption Filter	VetEquip	931401	Delivery system safety measures.
Model 923-B Mouse Gas Anesthesia Head Holder	KOPF	923-B	For animal restriction and O2 and isoflurane delivery.
Oxygen regulator, E-cylinder	VetEquip	901305	For O2 regulation and delivery.
Oxygen hose – green	VetEquip	931503	For O2 and isoflurane delivery.
Infrared Sterilizer 800 ºC	Marconi	MA1201	For instrument sterilization.
Surgical Instruments
Fine Scissors - ToughCut	Fine Science Tools	14058-11	For incision.
Surgical Hooks	INJEX	1636	In House Fabricated - Used to clear the surgical site from skin and fur.
Standard Tweezers or Forceps	-	-	For skin grasping.
Surgical Consumables
Vetbond	3M	SC-361931	For incision closing.
Cement and Catalyzer KIT (Duralay)	Reliance	2OZ	For implant fixation.
Sterile Cotton Swabs (Autoclaved)	JnJ	75U	For surgical site antisepsis.
24 Well Plate (Tissue Culture Plate)	SARSTEDT	831,836	For cement preparation.
Application Brush	parkell	S286	For cement mixing and application.
Pharmaceutics
Xylazin (ANASEDAN 2%)	Ceva Pharmaceutical (Brazil)	P10160	For anesthesia induction.
Ketamine (DOPALEN 10%)	Ceva Pharmaceutical (Brazil)	P30101	For anesthesia induction.
Isoflurane (100%)	Cristália (Brazil)	100ML	For anesthesia maintenance.
Lidocaine (XYLESTESIN 5%)	Cristal Pharma	-	For post-surgical care.
Ketoprofen (PROFENID 100 mg)	Sanofi Aventis	20ML	For post-surgical care.
Ringer's Lactate Solution	SANOBIOL LAB	7898153652145	For post-surgical care.
TobraDex (Dexamethasone 1 mg/g)	Alcon	631	For eye lubrification and protection.
Stimulation
Animal Transcranial Stimulator	Soterix Medical	2100	For current generation.
Pin-type electrode Holder (Cylindrical Holder Base)	Soterix Medical	2100	Electrode support (Implant).
Pin-type electrode (Ag/AgCl)	Soterix Medical	2100	For current delivery (electrode).
Pin-type electrode cap	Soterix Medical	2100	For implant protection.
Body Electrode (Ag/AgCl Coated)	Soterix Medical	2100	For current delivery (electrode).
Saline Solution (0.9%)	FarmaX	7896902206441	Conducting medium for current delivery.
Standard Tweezers or Forceps	-	-	For tDCS setup.
Real Time Polymerase Chain Reaction
BioRad CFX96 Real Time System	BioRad	C1000	For qPCR
SsoAdvancedTM Universal SYBR Green Supermix (5 X 1mL)	BioRad	1725271	For qPCR
Hard Shell PCR Plates PCT COM 50 p/ CFX96	BioRad	HSP9601	For qPCR
Microseal "B" seal pct c/ 100	BioRad	MSB1001	For qPCR