This 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.
It is widely acknowledged that the use of general anesthetics can undermine the relevance of electrophysiological or microscopical data obtained from a living animal’s brain. Moreover, the lengthy recovery from anesthesia limits the frequency of repeated recording/imaging episodes in longitudinal studies. Hence, new methods that would allow stable recordings from non-anesthetized behaving mice are expected to advance the fields of cellular and cognitive neurosciences. Existing solutions range from mere physical restraint to more sophisticated approaches, such as linear and spherical treadmills used in combination with computer-generated virtual reality. Here, a novel method is described where a head-fixed mouse can move around an air-lifted mobile homecage and explore its environment under stress-free conditions. This method allows researchers to perform behavioral tests (e.g., learning, habituation or novel object recognition) simultaneously with two-photon microscopic imaging and/or patch-clamp recordings, all combined in a single experiment. This video-article describes the use of the awake animal head fixation device (mobile homecage), demonstrates the procedures of animal habituation, and exemplifies a number of possible applications of the method.
An exciting recent trend in the Neurosciences is to develop experimental approaches for molecular and cellular probing of neuronal networks in the brain of awake, behaving rodents. Such approaches hold promise to shed new light on neurophysiological processes that underlie motor function, sensorimotor integration, perception, learning, memory, as well as injury progression, neurodegeneration and genetic diseases. Furthermore, recording from awake animal’s brain holds promise in development of novel therapeutic agents and treatments.
There is a growing awareness that anesthesia, which has been commonly used in neurophysiological experiments, can affect the basic mechanisms of brain function, potentially leading to erroneous interpretation of experimental findings. Thus, the widely used anesthetic ketamine rapidly increases formation of new dendritic spines and enhances synaptic function1; another commonly used anesthetic isoflurane at surgical anesthesia levels completely suppresses spontaneous cortical activity in new-born rats and blocks spindle-burst oscillations in adult animals2. At present, only a limited number of approaches enable experiments in non-anesthetized mice by means of two-photon microscopic imaging or patch-clamp recordings. These approaches can be divided into freely moving and head-fixed preparations.
The unique attractiveness of a freely moving animal preparation is that it allows assessment of natural behavior, including whole body movements during navigation. One way to image inside the brain of a freely moving rodent is to attach a miniaturized head-mounted microscope or fiberscope3-5. However, miniaturized devices tend to have limited optical performance in comparison to objective-based two-photon microscopy, and cannot be easily combined with whole cell patch-clamp recordings6.
The existing solutions for head-fixing an awake rodent have relied primarily either on physical restraint7,8 or on training the animal to exhibit voluntary head restraint9. Another popular approach is to allow the animal’s limbs to move by placing it on, e.g., a spherical treadmill10; this approach is often combined with computer-generated virtual reality. Electrophysiological experiments on head-fixed mice have mostly used extracellular recordings and were used to study central regulation of cardiovascular function11, effects of anesthesia on neuronal activity12, the auditory response in the brainstem13 and information processing14. The pioneering intracellular/whole-cell recordings in awake behaving animals were performed in the 2000s and have focused on neural activity related to perception and motion15-20. Around the same time, the first microscopic imaging studies on awake mice were published, where two-photon microscopy was used in the sensory cortex of physically restrained rats7 and on mice running on a spherical treadmill21.
Subsequent in vivo microscopy and electrophysiology studies demonstrated that a head fixation preparation can be successfully combined with behavioral paradigms based on forelimb movements, odor recognition, whisking, and licking8,22-25. Mice placed on the spherical treadmill can be trained to navigate the virtual visual environment generated by a computer10,26. Intracellular/extracellular recordings demonstrated that, in a head-fixed animal navigating such virtual environment, activation of hippocampal place cells can be detected27. In a virtual visual environment, mice demonstrate normal movement-related theta rhythm in the local field potential and theta-phase precession during active movement27. Recently, the spatial and temporal activity patterns of neuronal populations were recorded optically in mice during working memory decision tasks in a virtual environment28.
Despite having enabled breakthrough research, the spherical treadmill design has several inherent limitations. First, the animal is required to move on an unlimited surface of a rotating air-lifted ball, which poses no tangible obstacles such as walls or barriers. This limitation is only in part compensated by the computer-generated “virtual reality”, because visual input is arguably less effective on mice and rats as compared to the tactile sensory input (e.g., whisker-touch or lick), which these species naturally rely on. Second, the considerable curvature of the ball surface may be uncomfortable for the laboratory mice used to walking on a flat floor in their cages. Finally, the sheer diameter of the ball (at least 200 mm for mice and 300 mm for rats) renders the vertical size of the spherical treadmill device relatively large. This makes it difficult to combine spherical treadmill with the majority of commercially available microscopy setups, and often requires building a new setup around the treadmill by means of custom-made microscope frames.
Here, a novel method is described where a head-fixed mouse can move around an air-lifted mobile homecage that features a flat floor and tangible walls, and explore the physical environment under stress-free conditions. This article demonstrates the procedures of mouse training and head fixation, and provides representative examples where two-photon microscopy, intrinsic optical imaging and patch-clamp recordings are performed in the brain of awake behaving mice.
All the procedures presented here were performed according to local guidance for animal care (The Finnish Act on Animal Experimentation (62/2006)). The animal license (ESAVI/2857/04.10.03/2012) was obtained from local authority (ELÄINKOELAUTAKUNTA-ELLA). Adult mice (age 1-3 months, weight 20-40 g) were kept in group-housing cages in the certified animal facility of the University of Helsinki and provided with food and water ad libitum.
1. Cranial Window Implantation
Cranial window is implanted according to published protocols 29-31 with minor modifications, as briefly described below:
2. Animal Handling
3. Animal Training
NOTE: During prolonged training sessions that last more than 1-2 hr, consider providing the mouse with drinking water, which can be delivered either manually or using a pipette holder attached to the mobile homecage frame. Alternatively, water can be supplied for ad libitum use of the animal by placing viscous drops of hydro-gel directly on the walls of the mobile homecage.
NOTE: Remember to weigh the animals every day before training to rule out any chronic stress effects. Exclude an animal from the experiment if, at any time point, it demonstrates stress reactions such as freezing, vocalization, or stress-induced diarrhea.
4. Applications
The method presented here is intended for microscopic imaging or single-cell electrophysiological recordings in awake, head-fixed but otherwise freely-moving and behaving mice. The animal can move in two dimensions in a real (as opposed to virtual), tangible and familiar environment, while the animals’ skull is fixed firmly to the head fixation arm. Habituating the mice to the air-lifted mobile homecage consists of 4-6 days of twice-daily 2-hr training sessions (Figure 1). The trained animals can then be used in the experiments immediately. A typical study includes a number of imaging sessions or patch-clamp recording sessions that are spaced at intervals ranging from few hours to several days or weeks. Importantly, both optical and electrophysiological recordings can be performed simultaneously with cognitive or behavioral stimuli and readouts, within a single experiment.
To evaluate the mechanical stability of mouse’s head fixation in the mobile homecage, the image sequences of cortical vessels labeled with fluorescent-conjugated dextran and of cortical dendrites expressing YFP were collected while the experimental animals were navigating the mobile homecage (Figure 2). The maximal displacements of the brain during animal’s locomotion did not typically exceed 1-1.5 micrometers. These displacements occurred in the horizontal directions and very rarely resulted in a detectable shift of the imaging plane, rendering unnecessary any correction of motion artifacts. Stable head fixation in mobile homecage allows quantification of individual dendritic spines in awake animals with the same reliability as in anesthetized mice. Dendritic spine density, morphology and turnover can be monitored during longitudinal studies with multiple imaging sessions performed at intervals ranging from a few hours to several days or weeks.
The usability of the mobile homecage for functional optical imaging was tested in somatosensory cortex of awake mice using two approaches: i) two-photon microscopy on the Thy1-GCaMP3 transgenic mice and ii) intrinsic optical signal imaging in wild-type mice. Ca2+ imaging was performed in layer 2/3, which contains cell bodies of many fluorescently labeled neurons, as well as their dendrites and axons (Figure 3). The plots of fluorescence-over-time from selected regions of interest (ROIs) are shown in Figure 3, demonstrating spontaneous neuronal activity (measured as transient increases in GCaMP3 fluorescence) during the mouse’s active navigation in the mobile homecage. Optical imaging based on intrinsic signals allows mapping the spatial distribution of functional domains. Figure 4 illustrates wave-like changes in the blood oxygenation level (reflecting regional neuronal activation) that propagated along somatosensory cortex in response to vibrissa stimulation at the frequency of 0.05 Hz.
To test feasibility of patch-clamp recordings with mobile homecage, we used 2-3 months-old C57Bl/6J mice. Layer 2/3 neurons in somatosensory cortex were recorded from in whole cell configuration using current clamp mode. Patch-clamp recording in the brain of awake mice head-fixed on mobile homecage was essentially similar to blind patch-clamping in brain slices. Approximately 50% of attempts resulted in successful gigaseal formation, of which more than 70% yielded stable whole-cell configuration recording. No events of losing gigaseal contact due to mechanical displacement of cells were observed. Figure 5 illustrates a 60-sec fragment of a representative 10-min long current-clamp recording correlated with episodes of mouse’s active (running) and passive (resting) states.
Figure 1. The method of head fixation of awake mice in the mobile homecage. A) Overview of the air-lifted mobile homecage design and illustrations of the general concept. B) Diagram of a typical experimental timeline. The study begins with implantation of the cranial window two weeks prior to habituating the mouse to handling and wrapping, which is followed by eight twice-daily training sessions. The typical study includes a number of imaging sessions or patch clamp recording sessions that are spaced at intervals ranging from few hours to several days or weeks. Both optical and electrophysiological measurements can be done in parallel with cognitive or behavioral stimuli and readouts within a single experiment. Please click here to view a larger version of this figure.
Figure 2. Example of two-photon microscopic imaging on awake mice moving around the mobile homecage. A,B) Cortical vasculature, labelled with the 70 kDa Texas Red-conjugated dextran. The diameter of individual vessel segments is measured by plotting over time the profile of the lines drawn across the vessel lumen during periods of mouse’s resting and running (A). The rate of blood flow in arteries and veins is measured by line-scanning along the lines drawn parallel to the vessel wall (B). C,D) Fine details of neuronal morphology visualized in the brain of transgenic mice that express YFP in subpopulation of neurons under the Thy1 promoter. Three-dimensional reconstruction of pyramidal neurons in the mouse’s somatosensory cortex (C). The images of a dendritic branch acquired in an awake, behaving mouse are sufficiently stable for quantification of individual dendritic spine morphology (D). E) Quantification of brain motion caused by mouse movements. Larger amplitude displacements correlate with periods of the mouse’s running. Please click here to view a larger version of this figure.
Figure 3. Example of neuronal population activity in awake Thy1-GCaMP3 mouse moving around the mobile homecage. A) Two-photon image of cortical layer II/III neurons. ROIs, for example neuronal cell bodies, dendrites and axons are shown in yellow. B) ΔF/F traces of the GCaMP3 fluorescence from ROIs shown in A (time series recorded at 1.5 sec/frame). C) Zoomed-in region imaged at 65 msec/frame. D) Fluorescence from yellow ROIs in C plotted over time, shows the transient increases (red) in the GCaMP3 fluorescence that correspond to the action potential-induced Ca2+ influx episodes. Please click here to view a larger version of this figure.
Figure 4. Example of mapping the spatial distribution of functional responses in the cortex of an awake mouse by means of imaging the intrinsic optical signals. A) Bright-field view of the superficial blood vessels through the cranial window. B) Magnitude map of the baseline activity in mobile homecage during a 6-min episode. C) Magnitude map of neuronal activity propagating along somatosensory cortex in response to the vibrissa stimulation at a frequency of 0.05 Hz. Please click here to view a larger version of this figure.
Figure 5. Example of whole-cell patch-clamp recording in the cortex of an awake mouse moving around the mobile homecage. A) Current-clamp recording from a neuron in the mouse cortical layer 2/3. A 0.5 sec, 100-pA current injection (indicated below the trace) results in a burst of action potentials. The cell showed spike frequency adaptation characteristic for pyramidal neurons. B) Continuous current-clamp recording from the same neuron correlated with the mouse’ locomotor activity (shown in pink above the trace). Representative spontaneous activity of the layer 2/3 neuron during periods of the mouse’s resting (C) and running (D). Please click here to view a larger version of this figure.
Figure 6. Animal weight loss and locomotor activity of head-fixed/non-fixed mice during training sessions in the mobile homecage. A) Animal weight (mean+SD,%) before training sessions. Note that weight loss is fully reversed by the 7-8th training session. B) Trajectory of the mouse’s horizontal locomotion relative to the mobile homecage, which was extrapolated from the tracked movement of the mobile homecage during 8th training session. C) Tracked movements of a non-head-fixed mouse exploring the round cage during 8th training session. D) Duration of head-fixed (circle) and non-fixed (triangle) mice movement during 1-4th day of training (mean+SD,%). Note that, on day 4, head-fixed mice display neither freezing (as on day 1) nor excessive locomotor activity. Please click here to view a larger version of this figure.
To better understand brain physiology and pathology, research must be performed on a variety of preparation complexity levels, utilizing the most appropriate techniques for each preparation. At present, a wide range of neuroscience methodologies (from full-body fMRI to sub-organelle STED microscopy) are readily applied to anaesthetized animals, while experiments on awake and behaving animals have represented a significant methodological challenge.
Here, a novel approach is described where a laboratory animal, despite being firmly head-fixed, can move around an air-lifted mobile homecage and explore its tangible environment under stress-free conditions. The head-fixed behaving animal preparation presented here provides a number of crucial advantages. First, electrophysiological or imaging data obtained with this method are uncompromised neither by anesthesia nor by constrain-induced stress. Positioning of the mouse into the mobile homecage is quick and does not require anesthetizing the animal even transiently. Second, the air-lifted homecage ensures the mechanical stability that is needed to quantify changes in fine neuronal morphology and to record single-cell electrophysiological activity in awake animals. Finally, the mobile homecage’s design is more compact in comparison to the spherical treadmill, thus allowing positioning the mobile homecage under a standard upright microscope for two-photon imaging or patch-clamp recording in awake mouse’s brain.
Firm head fixation in the mobile homecage requires implantation of a specially designed four-winged metal holder, with a round opening in the center for optical or electrical access to the underlying brain region. These metal holders are attached to the skull by means of a combination of glue, dental cement and a small bolt screwed into the skull bone. This surgical procedure was developed based on a large number of previously published procedures, and was found to result in a stable and reproducible cranial window preparation. For in vivo electrophysiological experiments, a moon-shaped window34, a small size craniotomy (less than 0.5 mm)32, and a drilled glass-covered preparation35 have been utilized. Here, the “inverted” cranial window was implanted with either a large (3.5 mm diameter) or small (less than 0.5 mm diameter) craniotomy. Minimizing brain movement is critical for stable single cell recordings, which is why it is advisable to perform small size craniotomies for electrophysiological experiments. Upon implantation of the cranial window for optical imaging experiments, the animals are allowed to recover for at least 2 or 3 weeks, during which period the window first transiently loses its transparency and then regains it (with a 50-70% yield, depending on the genetic background of the mouse strain). Transparency of the cranial window and stability of the dental cement “cap” attached to the skull can be verified by means of a regular binocular microscope and physical inspection during animal handling. At the end of the 2-3 week recovery period, those animals that exhibit any signs of residual post-operational inflammation or mechanical defects in the dental cement should be excluded from the experiments and terminated.
The optimal age for starting training the mice is 2-4 months (corresponding to the body weight of 20-40 g). In younger animals, anchoring of the dental cement “cap” to the skull can be unreliable, which may decrease its resilience to the mechanical stress that is imposed by locomotion of the head-fixed mouse in the mobile homecage. Although male and female mice appear equally willing to navigate in mobile homecage, there is a tendency to achieve better percentage of cranial windows regaining their transparency in female mice (data not shown). Hence, in order to ensure a balanced mix of genders in the cohort of animals selected for imaging, implanting cranial windows in approximately 30% more male mice is recommended. Social interactions are known to improve the animals’ well-being and reduce stress, therefore it is advisable that littermates are operated and trained in parallel and kept together in group-housing cages.
In contrast to the procedures published for the spherical treadmill preparation13, the method utilizing the mobile homecage does not require anesthetizing the mouse at the moment of head fixation. This difference is important because it allows to rule out any residual effects that even a brief and “light” anesthesia episode is likely to have on the physiological measurements obtained shortly after. Indeed, even though in the studies where head fixation was done under anesthesia and the actual experiments were started after a brief waiting period13, one cannot exclude possible long-lasting effects of the brief anesthesia episode on the experimental data. Other studies have relied on water deprivation for systematic habituation of the animals to head fixation and used water reward as the means of motivating the animal to remain immobile36. However, the reward-based head fixation method limits the choice of applicable behavioral tests and, importantly, occupies one of the well-established stimulus–reward associations. In contrast, the method of mouse habituation to head fixation in mobile homecage does not require water deprivation and subsequent reward.
Supplementing the mobile homecage with a water delivery system is recommended for long-lasting experiments. The animal training sessions and experiments presented here were done during daytime (between 8 a.m. and 6 p.m.), which corresponds to the physiologically passive period for those mice that are kept under the standard 12-hr light schedule (lights on at 6 a.m. and off at 6 p.m.). Since the water intake is directly associated with the mouse’s activity, during the passive period mice do not require water delivery if the duration of a training/imaging/recording session does not exceed 2 hr. In addition to the timing and duration of the training sessions, one needs to address the issue of the optimal number of sessions required for habituating the animals to mobile homecage. To this end, two criteria were used to evaluate stress induced by head fixation procedures: i) weight loss, and ii) locomotor activity level. As shown in Figure 6, weight loss reaches the average level of 6% on training day 2, and is completely reversed by training day 4 (Figure 6A). Consistently with the weigh dynamics, the locomotor activity level of head-fixed animals is suppressed on the first day of training but stabilizes by training day 4 (Figure 6D). Based on these measurements, we suggest that the minimal duration of the mouse training period on mobile homecage is 4 days, as described in the protocol hereby.
Use of the air-lifted, flat-floored mobile homecage allows adding complex tasks (sensorimotor, perceptional, and cognitive) to the training paradigms for head-fixed mice. In the present study two protocols of behavioral tests are presented. Both protocols utilize odor cues and can be combined with longitudinal imaging/recordings in the mouse cortex. Although the mobile homecage is manufactured from nonabsorbent materials, one still needs to take into account possible interferences between the smell of the device and test odor(s). Another factor that may interfere with visual/tactile cues of a behavioral experiment is the junction between the wall and the insert, which is not seamless and may, therefore, be perceived by the animal as a landmark. It is worth noticing here that, in order to minimize animal’s distress during such interventions as placement of an odor-presenting cotton to the mobile homecage wall, the experimentalist should practice to perform such interventions as quickly as possible and avoid prolonged handling of the carbon cage. Alternative strategies for novel smell/object presentation are conceivable, e.g., placing hydrogel-based solution drops or objects (such as food chips) onto small shelves attached to the inner surface of the carbon cage wall at the height compatible with the animal’s head positioning.
Mobile homecage allows head-fixed animals to perform a wide range of two-dimensional movements including horizontal locomotion, situp, grooming, whisking, licking, nose-poking, skilled front paw movements, and wall touching with forelimbs, as illustrated in the present study. Using mobile homecage and the protocols presented here, researchers can study the sensorimotor neuronal system with a high level of control over both the stimulation conditions and the behavioral read-outs. Furthermore, studies of cognitive abilities in awake mice can be performed during conditioning, spatial navigation and decision-making tasks.
There are several practical limitations of this method. First, a significant amount of pressurized air is needed to achieve the homecage-lifting power and to perform long-lasting experiments. Second, the mobile homecage in its present implementation is only 18 cm in diameter, and therefore provides a relatively small and simple space in comparison to virtual reality, where a complex experimental environment can be designed without any spatial restrictions. Third, during whisker stimulation and reward-based experiments presented here, a device was used that limits the possibility the wall-contact for the mouse. Addition of an external visual or sensory stimulation channel (such as an eye-directed light projector) would require designing a more ergonomic and compact device in comparison to the multiple-screen or dome-projection solutions that have been used in the spherical treadmill experiments.
In summary, the use of the head-fixed mice moving in the air-lifted mobile homecage greatly facilitates the studies that combine cellular, molecular and behavioral levels of observation and manipulation within a single experiment. Specific applications illustrated here include two-photon microscopic imaging, intrinsic optical signal imaging and patch-clamp recordings in non-anesthetized behaving mice. It is expected that this approach will open new horizons in experimentation on awake, behaving mouse and serve as a useful tool for both drug development and basic research of brain function.
The authors have nothing to disclose.
The authors thank Prof. Eero Castren for his valuable comments on the manuscript. The work is supported by grants from The Academy of Finland, Centre for International Mobility of Finland, and Finnish Graduate School of Neuroscience (Brain and Mind Doctoral Program).
Name of Material/ Equipment | Company | Catalog Number | Comments/Description |
Tweezers Stainless Steel, 115mm | XYtronic | XY-2A-SA | |
Animal trimmer, shaving machine | Aesculap | Isis GT420 | |
Binocular Microscope | Zeiss | Stemi 2000 | |
Biological Temperature Controller with stainless steel heating pad | Supertech | TMP-5b | |
Blunt microsurgical blade | BD | REF 374769 | |
Borosilicate tube with filament | Sutter Instruments | BF120-69-10 | For patch pipette production |
Camera | Foscam | FI8903W | Night visibility |
Carprofen | Pfizer | Rimadyl vet | |
Dental cement | DrguDent, Dentsply | REF 640 200 271 | |
Dexamethasone | FaunaPharma | Rapidexon vet | |
Disposable drills | Meisinger | HP 310104001001008 | |
Dulbeco’s PBS 10X | Sigma | D1408 | |
Dumont #5 forceps, 110 mm | FST | 91150-20 | |
Eyes-lubricant | Novartis | Viscotears | For eyes protection during operation and as viscose solution for immersion |
Foredom drill control | Foredom | FM3545 | |
Foredom micro motor handpiece | Foredom | MH-145 | |
Four-winged metal holder | Neurotar | ||
Head Holder for Mice | Narishige | SG-4N | Assembled on stereotaxic instrument |
Hemostasis Collagen Sponge | Avitene, Ultrafoam BARD | Ref 1050050 | |
Imaris | Bitplane | ||
Ketamine | Intervet | Ketaminol vet | |
Kwik-Sil | WPI | ||
Mai Tai DeepSee laser | Spectra-Physics | ||
Micro dressing forceps, 105 mm | Aesculap | BD302R | |
Microelectrode puller | Narishige | PC-10H | Vertical puller for glass pipette production |
Micromanipulator | Sensapex | ||
Mini bolt | Centrostyle | Ref. 00343 s/steel M1.0x4.5 | |
Mobile Homecage | Neurotar | ||
Multiphoton Laser Scanning Microscope | Olympus | FV1000MPE | |
Nonwoven swabs 5×5 | Molnlycke Health Care | Mesoft | Surgical tampons |
Polyacrylic glue | Henkel | Loctite 401 | |
Round glass coverslip | Electron Microscopy Sciences | ||
1.5 thickness | |||
Small animal stereotaxic instrument | David Kopf Instruments | 900 | |
Student iris scissors, straight 11.5 cm | FST | 91460-11 | |
Xylazine | Bayer Health Care | Rompun vet |