In Vivo Wireless Optogenetic Control of Skilled Motor Behavior

Summary

The present protocol describes how to use wireless optogenetics combined with high-speed videography in a single pellet reach-to-grasp task to characterize the neural circuits involved in the performance of skilled motor behavior in freely moving mice.

Abstract

Fine motor skills are essential in everyday life and can be compromised in several nervous system disorders. The acquisition and performance of these tasks require sensory-motor integration and involve precise control of bilateral brain circuits. Implementing unimanual behavioral paradigms in animal models will improve the understanding of the contribution of brain structures, like the striatum, to complex motor behavior as it allows manipulation and recording of neural activity of specific nuclei in control conditions and disease during the performance of the task.

Since its creation, optogenetics has been a dominant tool for interrogating the brain by enabling selective and targeted activation or inhibition of neuronal populations. The combination of optogenetics with behavioral assays sheds light on the underlying mechanisms of specific brain functions. Wireless head-mounted systems with miniaturized light-emitting diodes (LEDs) allow remote optogenetic control in an entirely free-moving animal. This avoids the limitations of a wired system being less restrictive for animals' behavior without compromising light emission efficiency. The current protocol combines a wireless optogenetics approach with high-speed videography in a unimanual dexterity task to dissect the contribution of specific neuronal populations to fine motor behavior.

Introduction

Motor skilled behavior is present during most movements performed by us, and it is known to be affected in several brain disorders^1,2,3,4,5,6. Implementing tasks that allow studying the development, learning, and performance of skilled movements is crucial to understanding the motor function's neurobiological underpinnings, especially in models of brain injury, neurodegenerative and neurodevelopmental disorders^{2,7,8,9,10,11,12,13}. Reaching for and retrieving objects is done routinely in everyday life actions, and it is one of the first motor skills acquired during early development and then refined through the years^5,6. It comprises a complex behavior that requires sensory-motor processes such as the perception of the object's features, movement planning, action selection, movement execution, body coordination, and speed modulation^7,14,15,16. Thus, unimanual high dexterity tasks require the participation of many brain structures of both hemispheres^{16,17,18,19,20,21,22}. In mice, the single pellet reach-to-grasp task is characterized for several phases that can be controlled and analyzed separately^7,13,23. This feature allows to study the contribution of specific neuronal subpopulations at different stages of acquisition and behavior performance and provides a platform for detailed studies of motor systems^13,23,24. The movement occurs in a couple of seconds; thus, high-speed videography should be used for kinematic analysis in distinct stages of the skilled motor trajectory^7,25. Several parameters can be extracted from the videos, including body posture, trajectory, velocity, and type of errors²⁵. Kinematic analysis can be used to detect subtle changes during wireless optogenetic manipulation^7,23.

Using miniaturized light-emitting diodes (LEDs) to deliver light via a wireless head-mounted system makes it possible to have remote optogenetic control while the animal performs the task. The wireless optogenetic controller accepts single-pulse or continuous trigger commands from a stimulator and sends infrared (IR) signals to a receiver connected to the miniaturized LED^23,26. The current protocol combines this wireless optogenetics approach with high-speed videography of a dexterity task to dissect the role of specific neuronal populations during the performance of fine motor behavior²³. Since it is a unimanual task, it allows for assessing the participation of structures in both hemispheres. Traditionally, the brain controls the body movement in a highly asymmetric manner; however, high dexterity tasks require careful coordination and control from many brain structures, including ipsilateral nuclei and differential contribution of neuronal subpopulations within nuclei^{10,20,21,22,23}. This protocol shows that subcortical structures from both hemispheres control the trajectory of the forelimb²³. This paradigm can be suitable to study other brain regions and models of brain disease.

Protocol

The procedures involving animal use were conducted following local and national guidelines and approved by the corresponding Institutional Animal Care and Use Committee (Institute of Cellular Physiology IACUC protocol VLH151-19). Drd1-Cre transgenic male mice²⁷, 35-40 days postnatal with C57BL/6 background were used in the current protocol. Mice were kept under the following conditions: temperature 22±1 °C; humidity 55%; light schedule 12/12 h with lights off at 7 p.m. and were weaned at postnatal day 21. Weaned pups were housed in same-sex groups of 2-5. Animals were housed in static housing with micro-barrier tops. Bedding consisted of sterile aspen shavings. Rodent pellets and RO-purified water were provided ad libitum, except when noted.

1. Surgical procedures

1. Prepare an LED cannula at the desired length according to the dorsoventral coordinates of the structure of interest (ideally 0.5 mm longer to account for the thickness of the skull, for the dorsolateral striatum 3.5 mm) (Figure 1).
   1. Cut the glass fiber to a length longer than the final desired size, grind the fiber tip to the target length with rough sandpaper, and finally, polish the fiber tip with fine sandpaper.
      NOTE: LED cannula is a glass optical fiber of 250 µm diameter attached to an infrared receiver (see Table of Materials).
2. Pull glass pipettes (1.14 mm outer diameter, 0.53 mm inner diameter, and 3.5 in length) for the nano-injector with a horizontal puller (see Table of Materials) and store them for later. Program the puller in one loop to get a 15-20 µm tip diameter with a long gradual slope taper (4-5 mm).
3. Prepare the surgery area by thoroughly disinfecting the stereotaxic apparatus, hood, micro-injector (see Table of Materials), and surrounding surfaces with 70% ethanol.
   NOTE: A mouse stereotaxic apparatus is essential to inject Adeno Associated Virus (AAVs) precisely and place the LED cannula in the region of interest.
4. Wear the appropriate personal protective equipment for the procedure, including a clean lab coat or disposable surgical gown, sterile gloves, face mask, and disposable head cap.
5. Place the necessary equipment close to the surgery area, such as sterile surgical tools, cotton tips, solutions, micropipette, pipette tips, capillaries, micro-fill with mineral oil, and marker.
6. Fill a pipette for microinjections with mineral oil and place it in the micro-injector. Make sure that the micro-injector is working correctly by ejecting some mineral oil.
   NOTE: All instruments used during the surgery should be autoclaved and sterile. Aseptic technique should be used.
7. Anesthetize animals with gaseous isoflurane 4-5% to induce anesthesia and 1.2% throughout the surgery with 0.5-1 L/min pure oxygen. The surgery begins only after the animal has reached a point of deep anesthesia, assessed by the absence of paw withdrawal after a slight pinch.
   1. Continuously monitor the breathing rate and temperature of the animal. Maintain the body temperature by a heating pad set at 34 °C.
8. Apply an ophthalmic ointment. Remove hair from the scalp with a trimer and hair removal cream. Wipe the scalp with cotton swabs having 8% povidone-iodine (see Table of Materials) and 70% ethanol alternated three times each.
9. Place the mouse in the stereotaxic apparatus and secure the head, ensuring that the skull is leveled in the mediolateral and anterior-posterior axes.
10. Make a 1 cm incision with a scalpel through the scalp at the level of the eyes along the sagittal axis. Retract the skin to expose the skull and clean the periosteum with cotton swabs.
11. Clean the cranium surface with saline solution and sterile cotton swabs. Resolve any bleeding at the surface using sterile absorbent eye spears (see Table of Materials) or similar sterile absorbent material.
12. Apply a drop of 2.5% hydrogen peroxide with a cotton swab and let it act for a few seconds to make the skull sutures visible and have a better reference. After a few seconds, clean thoroughly with a clean cotton swab.
13. With the glass pipette (15 µm final tip diameter), locate bregma and lambda to check that the skull is leveled in the anterior-posterior axis.
    NOTE: It is recommended to have a stereoscopic microscope or USB microscope to see the tip of the glass pipette. In case it is needed, adjust the height of the mouth holder to level the skull.
14. Move the capillary toward the selected anterior-posterior (AP) and medial-lateral (ML) coordinates (dorsolateral striatum AP 1.2 mm, ML 2.28 mm). Paint a reference point in the scalp above the selected coordinates with a sterile marker.
15. In the reference point, perform an ~1 mm diameter craniotomy applying gentle pressure to the skull with a sterile rotary tool or dental drill at a low to medium speed with a small round dental drill bit (see Table of Materials).
16. Load the capillary with 300-400 nL of Cre-dependent adeno-associated virus (AAV) such as AAV1-dflox-hChR-2-mCherry to express Channelrhodopsin or an AAV to express only the reporter protein (e.g., mCherry) as a control in the region of interest (see Table of Materials). Check that the tip is not clogged, then introduce the glass pipette in the brain at the desired dorso-ventral (DV) coordinates (dorsolateral striatum DV -3.35 mm).
    1. Inject 200 nL using an automatic injector at a rate of 23 nL/s. Wait for 10 min after finishing the injection, withdraw the glass pipette slowly to avoid spillage.
       NOTE: It is possible to use a 30 G needle to inject with the appropriate micro-injector.
17. Clean and dry any residues with cotton swabs.
18. Attach the sterile glass LED cannula to the stereotaxic arm and calibrate the coordinates using bregma as a reference. Insert the cannula very slowly (300 µm/min) to avoid tissue damage and place it 100 µm above the injection site.
19. Once the LED cannula is in place, add a drop (100 µL) of tissue adhesive at the edge of the craniotomy.
20. Prepare dental cement mixture (see Table of Materials) following the manufacturer's instructions to fix the fiber to the skull.
    NOTE: Briefly, use a chilled porcelain dish to have more working time before cement sets. Add 2 scoops of resin clear powder to the porcelain dish, add 4 drops of quick base and 1 drop of catalyst, then mix well. The powder/liquid ratio can be adjusted if a thinner or thicker viscosity is needed.
21. Using a sterile brush, apply the dental cement mixture around the cannula connector little by little, building layers until the skull is covered and the connector is securely attached to the skull, leaving the pins completely free. Avoid getting dental cement on the skin of the mouse.
22. Allow drying completely.
23. Close the skin around the implant using tissue adhesive (see Table of Materials).
24. Place the mouse in a recovery cage over a heating pad at 33° C. Monitor for the presence of one or more of the following signs of pain/discomfort: 1) Hunched up, lack or reduction of motor activity, 2) Failure to groom reflected in a unkept dirty coat, 3) Excessive licking or scratching, redness in the incision site, 4) aggressive behavior, 5) anorexia or dehydration, and 6) Lack of nest formation.
    NOTE: Keep the mouse individually caged during all the procedures to avoid implant detaching. In case of detachment of the cannula, perform euthanasia by injecting 150 mg/kg of sodium pentobarbital followed by decapitation after deep anesthesia is reached.
25. Inject subcutaneously (SC) meloxicam 1 mg/kg once daily for three days post-surgery to provide analgesia.
26. Wait at least 7 days for complete recovery and 14 days for opsin expression before further procedures.
    NOTE: Perform a postoperative follow-up every 12 hours for three days, then check animals every day until the day of euthanasia at the end of the experiment.

2. Reach-to-grasp training

1. On day 7 post-surgery, start the food deprivation protocol²⁸. Weigh mice for three consecutive days to determine their average ad libitum body weight. Then, schedule food restrictions so that the animals receive enough nutrients to maintain approximately 90% and not less than 85% of body weight.
   NOTE: This is achieved by providing 2.5-3 g of food daily. Monitor animals' weight daily and score for overall well-being observing animals' behavior and appearance, for example, coat and eyes appearance. Use the body condition scoring system from Reference²⁹.
2. During the pre-training, training, and testing periods, provide each mouse with 20 pellets (20 mg of dustless chocolate-flavored pellets) daily (see Table of Materials) (eaten during the task or after) besides the standard food pellets.
3. Three days before habituation, scatter 0.4 g/animal/day 20 mg of dustless chocolate-flavored pellets in their home cages, so mice get acquainted with the pellets that serve as a reward during the reach-to-grasp task.
4. Habituate mice by placing them 10 min in the testing chamber one day before pre-training with pellets scattered on the chamber floor (Figure 1A).
5. Allow food daily after training and testing. Keep a similar schedule every day.
6. On the first day of pre-training, place the mice in the reach-to-grasp chamber and observe from the front. Place the pellets in front of the chamber close to the opening so that they start consuming the pellets. At this stage, mice are allowed to grab the pellets in any form.
7. On day two of pre-training, place the pellets further and further from the opening until getting them to the indentation (1 cm from the opening) so mice can shape their reach-to-grasp movement (Figure 1C).
8. Train mice to run to the rear of the cage and return to the cage opening to receive the next food pellet as a strategy to individualize trials.
   NOTE: This can be achieved by waiting until the mouse is in the rear of the cage before placing a pellet in the indentation for each trial.
9. Place pellets to be grasped by either their right or left paw.
   NOTE: Mice start using preferentially one paw to grasp, which will be used the following days of training and testing.
10. Train animals for 6 days in daily sessions lasting 20 trials or until a maximum of 10 min elapse. From day 2 of training, put the mock receiver (dimensions 12 x 18 x 7 mm, 1 g, see Table of Materials), so mice get habituated to the weight while performing the task (Figure 1B). Each day score the number of hit and missed trials.
11. Record behavior with a regular camera and capture 30-60 frames/s from the front of the chamber. Additionally, one can place a mirror under the training chamber at a 45° angle to monitor the animals' posture (Figure 1D,E).
12. For post-hoc kinematic analysis (Figure 2), mount a high-speed camera (see Table of Materials) at an angle of 45° to record from the side of the cage. If a 3D analysis is required, place a second high-speed camera to record at a 35° angle from the front of the chamber; both cameras should be placed in the right or left side of the cage depending on animals' sidedness and should capture at the same frame rate and be synchronized⁷ (Figure 3D,E).
13. Set the high-speed cameras to 100 frames/s with a resolution of 376 x 252 pixels or more if possible. Place white Styrofoam walls behind the sides and back of the chamber to reduce background and increase contrast (Figure 1E).
14. On test day, replace the mock unit with an infrared receiver for wireless optogenetic stimulation (Figure 1B,C).
15. When mice start reaching, turn the LED cannula manually with the remote controller to have a continuous stimulation for the time the behavior is performed and for no longer than 2 s. Programming an automatic stimulation paradigm is preferable. The stimulation device triggers an LED of 470 nm (blue light) with intensity at the tip of 1.0 mW/mm².
16. Collect the videos for further examination, including scoring and kinematic analysis.

3. Post-hoc histological confirmation

1. Upon completion of an experiment, confirm viral expression and LED cannula placement. Anesthetize the animal with a cocktail of ketamine 100 mg/kg and xylazine 10 mg/kg. Once the mouse presents signs of deep anesthesia (step 1.7), perfuse with ice-cold phosphate-buffered saline (PBS) followed by 4% PFA.
2. Remove implanted cannula carefully by firmly grasping the connector with forceps and pulling up gently.
3. Extract and post-fix the brain for 24 h in 4% PFA²³.
4. Perform 3-10 min washes with PBS.
5. Cut the brain in 50 µm sections using a microtome (see Table of Materials).
6. Mount the sections in slides with hard-set mounting media with DAPI to stain nuclei and cover slides.
7. After drying, observe the sections under the confocal microscope and verify the implanted cannula location and expression of Ch2R fused with any fluorescent protein.

Results

The reach-to-grasp task is a paradigm widely used to study shaping, learning, performance, and kinematics of fine skill movement under different experimental manipulations. Mice learn to execute the task in a couple of days and achieve more than 55% accuracy reaching a plateau after 5 days of training (Figure 2A,B). Similar to what has been previously reported, a percentage of animals do not perform the task appropriately (29.62%), and those should be excluded from further a...

Discussion

The use of optogenetic manipulation of neuronal populations in well-defined behavioral paradigms is advancing our knowledge about the mechanisms underlying motor control^7,23. Wireless methods are especially suitable for tasks that require tests on multiple animals or free movement^34,35. Nevertheless, as techniques and devices are refined, it should be the go-to option for any behavioral task combined with...

Materials

Name	Company	Catalog Number	Comments
Anaesthesia machine	RWD	R583S	Isoflurane vaporizer
Anesket	PiSA		Ketamine
Breadboard	Thorlabs	MB3090/M	Solid aluminum optical breadboard
Camera lense	Canon		50mmf/ 1.4 manual focus lenses (c-mount)
Camera system	BrainVision	MiCAM02	Camera controller and synchronizer
Cotton swabs
CS solution	PiSA		Sodium chloride solution 9%
Customized training chamber	In house
Drill bit #105	Dremel	2 615 010 5AE	Engraving cutter
Dustless precission chocolate pellets	Bio-Serv	F05301
Ethyl Alcohol	J.T.  Baker	9000-02	Ethanol
Eyespears	Ultracell	40400-8	Eyespears of absorbent PVA material
Fluriso	VetOne	V1 502017-250	Isoflurane
Glass capillaries	Drumond Scientific	3-000-203-G/X	Pipettes for NanoJect II
Hidrogen peroxide	Farmacom		Antiseptic
High-speed camera	BrainVision	MiCAM02-CMOS	Monochrome high-speed cameras
Infrared emmiter	Teleopto
Insulin syringe
LED cannula	Teleopto	TelC-c-l-d	LED cannula 250um 487nm light
Micropipette 10 uL	Eppendorf	Z740436
Micro-pipette puller	Sutter	P-87	Horizontal puller
Microscope LSM780	Zeiss		Confocal microscope
Microtome
Mock receiver	Teleopto
NanoJect II	Drumond Scientific	3-000-204	Micro injector
Oxygen tank	Infra	na
pAAV-EF1a-double.floxed-hChR2(H134R)-mCherry-WPRE- HGHpA	Addgene	20297	Viral vector for ChR-2 expression
Parafilm
Paraformaldehyde	Sigma	P-6148
Phosphate saline buffer	Sigma	P-4417	Phosphate saline buffer tablets
Pipette tips 10 uL	ThermoFisher	AM12635	0.5-10 uL  volume
Pisabental	PiSA		Sodium pentobarbital
Plexiglass	commercial		Acrylic sheet
Povidone iodine	Farmacom		Antiseptic
Procin	PiSA		Xylacine
Puralube	Perrigo pharma	1228112	Eye lubricant 15% mineral oil/85% petrolatum
Rotary tool	Kmoon	Mini grinder	Standard
Scalpel
Scalpel blade
Stereotaxic apparatus	Stoelting	51730D	Digital apparatus
Super-Bond C&B	Sun Medical		Dental cement
Surgical dispossable cap
Teleopto remote controller	Teleopto
Tg Drd1-Cre mouse line	Gensat	036916-UCD	Transgene insertion FK150Gsat
Tissue adhesive	3M Vetbond	1469SB
TPI Vibratome 1000 plus	Peico		Microtome
Vectashield mounting media with DAPI	Vector laboratories	H-1200	Mounting media
Wireless receiver	Teleopto	TELER-1-P