Hello, I'm Hel Lin, a post-grad student at the Near Science Center, university of Helsinki. Today my colleagues and I will show you how to perform optical and electrophysiological recordings in awake head fix, but otherwise free moving and behave animals. Our method creates a tangible, familiar environment for the mouse to navigate and explore.
During microscopic imaging or single cell electrophysiological recordings, which require firm fixation of the animal's head, we implant a chronic cranial window and habituate the mouse to an airlifted mobile home cage, which glides under the animal's feet. During locomotion on a flat surface, the mobile home cage can be positioned under a standard upright microscope for two photon imaging or placed into an electrophysiology set up for patch clamp recordings in a wake mouse brain. All experimental procedures presented here have been approved by the local ethics committee.
We implant cranial windows according to published protocols with minor modifications upon cranial window implantation. The animal is allowed to recover for at least two or three weeks during this recovery period, the window normally regains its transparency. Animals with any signs of residual post operational inflammation are excluded from the experiments.
Train the animals selected for a study to habituate them to head fixation. In the mobile home cage, handle the mouse calmly for five to 10 minutes, avoiding abrupt movements and noises, then return it to the home cage. Repeat the procedure two or three times at unequal intervals.
The mobile home cage allows head fixation without anesthetizing the animal prior to head fixation. Use a rag to wrap the mouse two or three times at unequal intervals. Make sure the animal remains calm while wrapped.
Otherwise, repeat the procedure until the mouse is fully habituated to it. Start the twice daily training of the mice on mobile home cage on the next day after the handling is completed. Weigh the animals before each training session.
Animals showing weight loss above 10%should be excluded from the study. Adjust the position of the head fixation arm to match the size of the trained animal. Connect the air inlet of the mobile home cage device to the standard laboratory pressurized air outlet.
Make sure that the air pressure is sufficient for free flotation of the airlifted home cage. Begin the training of habituated animals in the mobile home cage. Immobilize the mouse by wrapping it in the rag.
Insert the metal holder attached to the animal's head into the head fixation arm and firmly fix it by tightening the screws. Turn on the airflow to make the airlifted home cage levitate. Remove the rag and allow the mouse to explore the home cage.
During the first training session, mice may exhibit freezing and trembling behavior and tail erection. To habituate the animal to noise, provide a constant exposure to ambient sounds using, for example, a radio translation or recorded music and speech during all training sessions as well as during the experiments. To avoid chronic stress, use bright light illumination during the first hour of the training session.
After the first hour, conduct training in the dark and use a near infrared camera to monitor the animal's behavior. When the training session is completed, release the animal from the head fixation arm by losing the screws. Return the animal back to the home cage for at least two hours before the next training session.
Keeping the mouse in a cage with two or more other mice will decrease the stress level. Optionally, you can place the airlifted cage together with the animal in its home cage. This will enrich the environment and may reinforce the habitation of the animal to the mobile home cage.
During the last training session, mice are habituated to head fixation and should exhibit Stressless behavior navigating calmly around the airlifted home cage. In the absence of visible light, the trajectory of the mouse locomotion can be tracked by monitoring the cage movements with a near infrared camera or with an optical computer mouse. During prolonged training sessions that last more than one or two hours, consider providing the mouse with a drinking water, either manually or using a pipet holder attached to the mobile home cage frame.
The following diagram summarizes the timeline of a typical experimental study. The study begins with implantation of the cranial window two weeks prior to mouse handling, and continues with eight twice daily training sessions. The typical study includes a number of imaging sessions or patch clamp recording sessions that are spaced at an interval of a few hours to several days or weeks.
Both optical and electrophysiological experiments can be done in parallel with cognitive or behavioral stimuli and readouts. Two photon microscopy is a unique nonlinear optical technology that allows real time visualization of cellular and even subcellular structures in deep cortical layers of a rodents brain. To perform a two photon microscopic imaging experiment, place a trained animal in the mobile home cage, adjust the stoppers to protect the microscope optics from accidental damage.
Clean the cover glass of the cranial window to remove dust particles. Drop an immersion fluid on the cover glass. We recommend using viscus solution because water evaporates rapidly.
The platform can be placed on top of a typical motorized optical table using the wide field mode of the fluorescence microscope and a long pass emission filter. Assess the brain vasculature and select a region of interest. To image cortical vasculature inject into the tail vein of a head fixed and immobilized animal, a 70 kilo Dalton, Texas red conjugated text Tran.
This image sequence shows a time-lapse recording of venues and arterials in the cortex of a behaving mouse while it navigates around the mobile home cage. By plotting the profile of the lines drawn across the vessel lumen, one can measure the changes in the vessel diameter and the motility of individual vessel segments. The rate of blood flow in arteries and veins can be measured by line scanning along the lines drawn parallel to the vessel wall.Fine.
Details of neuron morphology can be visualized in transgenic mice that express YFP in the subpopulation of neurons under the Taiwan promoter. These two Zacks compare the imaging stability between anesthetized and awake animals. The right side panel shows dendrites from the animal anesthetized with ketamine xylazine mixture.
While the left side panel shows a dendrites image in an awake animal walking around a mobile home cage. Individual spine morphology in awake animals can be analyzed with the same reliability and precision as in the anesthetized mice. The displacement of the brain during animals running do not typically exceed one to one and a half micrometers.
These movements occur in the horizontal directions and very rarely result in a shift of the imaging plane making correction of motion artifacts unnecessary. In most cases, the ability to chronically monitor neuronal activity in the awake living brain is essential for understanding the organization and function of the nervous system. Using the tie one G Camp three transgenic mice, we image spontaneous neuronal activity in the awake mice behaving in the mobile home cage.
This time lapse image sequence shows representative examples of calcium spikes in neuronal cell bodies, nites and axons. Optical imaging based on intrinsic signals allows mapping the spatial distribution of functional domains to perform an intrinsic optical imaging experiment. Place the trained animal in the home cage positioned under the camera objective.
Clean the cover glass from dust. Put a drop of glycerol and cover it with a five millimeter round cover slip. Tune the position of a high speed camera.
Place a manipulator with a air blowing tube opposite to contralateral vibra. Use green lights to image the vessel map. Focus deeper into the cortex, approximately 400 micrometers below the cortical vessels.
Place a band pass filter in front of the camera and illuminate cortex with a red light. Allow the mouse to adopt to the platform for at least 30 minutes. Record the baseline activity during a six minute episode.
Stimulate vibra every 10 seconds with air puffs for the total period of six minutes. This image illustrates neural activity propagating along somatosensory cortex in response to vibra stimulation at the frequency of 0.05 hertz. To perform single cell electrophysiological recordings, place the trained animal implanted with an inverted cranial window in the mobile home cage.
Slowly and carefully, remove the cover glass from the metal holder. Refresh the cortical buffer and clean the cranial window from debris with a hemostatic tampon. Place the ground electrode into the cortical buffer.
Fill the patch clamp pipette with intracellular solution. Apply a positive pressure to the recording pipette using a motorized micro manipulator, deepen the pipette into the cortex and obtain a giga seal contact with the cell of interest. After recording, remove the pipette, refresh the cortical buffer and close the cranial window by glue in a cover glass to the metal holder.
The use of an airlifted home cage allows combining imaging and electrophysiology with classical behavioral paradigms such as learning, rehabilitation or anxiety related behavior. Olfaction is an important sensory input for mice and can be used for stimulation in memory tests, and for assessment of reaction to novelty. To perform a habituation disa olfactory test, place the trained animal in the mobile home cage and allow it to adopt.
For 30 minutes, attach a clean piece of cotton soaked in tap water to the inner wall of the mobile home cage using a two-sided adhesive tape. After five minutes, replace the clean cotton on the home cage wall with a new one soaked in a liquid containing the vanilla smell. Repeat the five minute exposures to the vanilla smell three times with five minute intervals between the sessions.
To measure the level of interest to the smell. Count the cumulative time that the animal spends facing the target zone and the total time of mobile home cage movement. During the last fifth session, present a socially significant smell.
Wet the new cotton applicator with a few drops of urine from mice of the opposite gender. To assess the reaction to novelty, divide the mobile home cage into four zones. Attach two cotton pieces opposite to each other.Color.
Code them with markers on the external side of the home cage walls during the first trial. Place two clean cotton pieces. Then replace both of them with cotton soaked in a 1%extract of vanilla.
During the third trial, put the vanilla cotton to zone number one and the piece of cotton soaked in a 1%banana extract to the zone. Number two, to measure the preference to no smell. Calculate the time that animals spend facing the no smell as percentage of the cumulative time facing other zones.
Hi, I'm Leo Kiro, a principal investigator at the Neuroscience Center University of Helsinki. I hope you've enjoyed watching the video and found it useful. We have been working with twofold on microscopy in vivo for the last five years, and the use of general and aesthetics has always been a great concern because an aesthetics interfere with the brain function, with the blood flow and everything.
And today I am happy that we can work in awake animals and perform stable microscopic imaging experiments and validate and verify the results that we had previously obtained in anesthetized animals. I'm sure that the use of mobile home cage and the methods that we presented today will increase the relevance of in vivo research in general and will lead to new and breakthrough results, and I'm looking forward to it.
