This work is supported by the National Natural Science Foundation of China (32271060). Y.-t.L. designed the research, performed the experiment, analyzed the data, and wrote the manuscript. Z.L. and R.W. performed the experiment.

All experimental procedures were performed in accordance with the animal welfare guidelines and approved by the IACUC at the Chinese Institute for Brain Research, Beijing.
NOTE: The timeline for this protocol is as follows: 1) make the suction cup; 2)&#160;inject the virus; 3) implant the headplate; 4)&#160;after 3 weeks, implant the plug; 5)&#160;after a ~3 day recovery and habituation on the treadmill, perform two-photon/wide-field imaging.
1. Preparation of a suction cup (Figure 1A)
<ol>
	<li>Deposit a drop of phosphate-buffered saline (PBS, 1x) in an acrylic dish and touch it with a 21 G flat needle to fill the tip by capillarity.</li>
	<li>Cover the tip of the needle with a translucent silicone adhesive and set for about ~10 min.</li>
	<li>Cut the silicone adhesive with scissors to a disk with a ~2 mm diameter.</li>
</ol>
2. Preparation of plugs (Figure 1B)
<ol>
	<li>Prepare a 0.75 mm thick plastic shim stock and cut out a 1&#160;cm x 1&#160;cm&#160;square in the center. Prepare two acrylic blocks. Clean the shim stock and acrylic blocks with alcohol and wipe them with lens paper.</li>
	<li>Put the shim stock on one acrylic block, deposit the silicone adhesive into the center to fill ~90% of the opening (avoid bubbles), add another acrylic block and ~1 kg force, and wait 20 min.</li>
	<li>Under a surgical microscope, cut the silicone adhesive with a scalpel to 1 mm high by 1.5 mm wide triangular prisms above a piece of paper with triangles printed on it.</li>
	<li>Remove any dust from the triangular prisms with plastic sticky tape and put it on a glass coverslip of 5 mm diameter and 0.15 mm thickness.</li>
	<li>Put the coverslip with the triangular prisms under a corona treater, turn on the corona treater, and bring it closer to the coverslip until lightening appears. Hold it at that position for a few minutes until they are attached to each other. 
	NOTE: This step might be different for other corona treaters.</li>
	<li>Put the plugs in a Petri dish and place it in an incubator at 70 &#176;C overnight.</li>
</ol>
3. Preparation of animals and injection of viral construct
<ol>
	<li>Use C57BL/6J (wild-type) and Emx1-Cre:Pals1flox/wt (partial-cortex)20 mice of both sexes at 6 weeks of age (9 weeks at the time of imaging).</li>
	<li>Sterilize all materials and instruments used for the surgical procedure.</li>
	<li>Anesthetize the mouse with 5% isoflurane and then maintain the isoflurane at 1%-2% with a flow rate of 1 L/min during the surgery. Confirm the level of anesthesia by the lack of toe pinch reflex.</li>
	<li>Head-fix the mouse on a stereotaxic frame with ear bars. Throughout the surgery, put ophthalmic ointment on the mouse eyes to prevent drying, and maintain its body temperature at 37 &#176;C with a thermostatic heating pad.</li>
	<li>Shave the fur and sterilize the skin surface with iodophor and 75% ethanol. Inject lidocaine (5 mg/kg, subcutaneous injection [SQ]), tilidine (2 mg/kg, SQ), and meloxicam (2 mg/kg, SQ). 
	NOTE: The surgical procedure for sterilization, analgesia, and local anesthesia may differ across institutions.</li>
	<li>Remove the skin over the SC with surgical scissors to expose the skull. Clean the skull with a cotton swab.</li>
	<li>Adjust the stereotaxic frame until the skull surface is flat. Drill a hole 0.5 mm lateral and 0.42 mm anterior to the lambda until the dura is exposed.</li>
	<li>Inject adeno-associated virus (AAV) expressing GCaMP6m (rAAV2/9-hsyn-GCaMP6m; 1 x 1013 GC/mL dissolved in 1x PBS) at depths of 1 mm and 1.6 mm from the lambda with a beveled glass micropipette (tip is beveled to 25&#176; with a diameter of 40-50 &#181;m).</li>
	<li>At each depth, inject 100 nL at a speed of 50 nL/min and wait 5 min after each injection. After the injection, slowly withdraw the injector.</li>
</ol>
4. Implantation of the headplate
<ol>
	<li>Clean the muscles and tissues over the skull and scratch the skull with a blade. If bleeding occurs, stop it with gel foam and clean the blood. Attach the headplate (Figure 1C) to the skull with a self-curing dental adhesive resin cement. Specifically, mix a 3/4 spoon of polymer, three drops of monomer, and one drop of catalyst in a ceramic bowl on ice. Apply the mixture on the headplate and the skull. Attach them together and wait 5 min for the adhesive to set.</li>
	<li>Inject antibiotics (ceftiofur sodium, 5 mg/kg, SQ) to prevent infections. Remove the mouse from the stereotaxic frame and place it on a heating pad for recovery. Return it to the home cage after recovering from anesthesia. For pain management, provide ibuprofen-containing drinking water&#160;(0.5 mL/50 mL) to the mouse for 3 days after the surgery.</li>
</ol>
5. Implantation of the plug (Figure 2)
<ol>
	<li>Implant the plug 3 weeks after the viral injection. Anesthetize and fix the mouse on a stereotaxic frame, as described in steps 3.2-3.5. Prepare gel foam soaked in saline to stop potential bleeding.</li>
	<li>Drill a 3 mm x 2 mm oval with a microdrill centered at 0.5 mm posterior to the lambda. Thin the boundary of the oval until it cracks. Thin the skull in front of the oval recording window to make the coverslip close to the SC.</li>
	<li>Take off bone flaps using fine forceps. Make a cut on the dura posterior to the transverse sinus and remove the piece of dura. If necessary, add artificial cerebrospinal fluid (ACSF) to the brain with a 3 mL syringe to prevent drying. Dry the skull and recording window with gel foam and cotton swabs.</li>
	<li>Deposit a drop of silicone adhesive into the recording window. Use the suction cup to hold the plug with negative pressure. Use a motorized micromanipulator to move the suction cup and lower the plug into the silicone adhesive until the coverslip touches the skull. Then, move the plug forward ~1 mm to push away the transverse sinus and expose the SC. This step should be done in 1-2 min before the silicone adhesive becomes sticky.</li>
	<li>Clean the headplate and apply butyl cyanoacrylate and resin cement around the boundary of the plug to fix it to the headplate. 
	NOTE: Provide postoperative care, as described in step 4.2.</li>
</ol>
6. Two-photon imaging (Figure 3)
<ol>
	<li>Head-fix the animal on a rotating treadmill with its headplate. Place the treadmill under a two-photon microscope and adjust the height to an appropriate position. Put a black aluminum cone between the objective and the headplate to prevent light contamination from the monitor for visual stimulation. Habituate the mouse for ~15 min.</li>
	<li>Perform two-photon imaging on the microscope with a 16x magnification, 0.8 numerical aperture (NA), and 3 mm working distance (WD) objective. Use a Ti:sapphire laser with mode-locking technique controlled by two galvo scanners to excite GCaMP6m at 920 nm. Adjust the laser power at the sample plane between 20-80 mW.</li>
	<li>Scan a 600 &#181;m x 600 &#181;m field of view at 4.8 Hz with a spatial resolution of 2.4 &#181; m/pixel, and image neural activity across the depth up to 350 &#181; m. 
	NOTE: The sampling rate and spatial resolution are for galvo scanners, and should be adjusted for resonant scanners.</li>
	<li>Collect emitted fluorescence with a dichroic mirror and detect it using a photomultiplier tube (PMT)&#160;after passing it through a bandpass filter.</li>
	<li>Record the mouse&#39;s locomotion on the treadmill with a rotary encoder. Use a camera with a 50 mm lens to record its pupil size and position. Synchronize these recordings and image acquisitions to visual stimulation by recording triggers sent by the stimulation computer.</li>
</ol>
7. Analysis of calcium responses measured by two-photon imaging
<ol>
	<li>Correct the brain motion during imaging using SIMA21 or NoRMCorre22.</li>
	<li>Manually draw a region of interest (ROI) using the Cell Magic Wand tool (ImageJ) and fit it with an ellipse in MATLAB. Extract fluorescence traces of each ROI after removing neuropil contamination: 
	<img alt="Equation 1" src="/files/ftp_upload/65181/65181eq01.jpg" style="margin: auto;" /> 
	Ftrue&#160;and Fraw are the corrected and raw fluorescence amplitudes of the ROI, respectively, Fneuropil is the fluorescence amplitude of the surrounding neuropil, and r is the out-of-focus neuropil contamination factor (0.7 for our setup). The r is estimated by measuring the signals on a blood vessel for which Ftrue = 0, as described previously23,24.</li>
	<li>Remove slow baseline fluctuations by subtracting the eighth percentile value from a 15 s window centered on each frame25.</li>
	<li>Define visual responses of a neuron as the relative change of fluorescence amplitude in its ROI during the stimulus period: 
	<img alt="Equation 2" src="/files/ftp_upload/65181/65181eq02.jpg" style="margin: auto;" /> 
	Fpeak&#160;is the peak amplitude of the fluorescence trace during the visual stimulation, and Fbaseline is the mean amplitude during a 0.5 s period prior to stimulation.</li>
</ol>
8. Wide-field imaging and data analysis (Figure 4)
<ol>
	<li>Build a wide-field macroscope with a tandem lens4 (Figure 4A). The tandem-lens macroscope is two camera lenses (50 mm and 105 mm) connected via an adapter, and the magnification is ~2x.</li>
	<li>Prepare partial-cortex mutant mice as described in steps 3.2-3.5. Inject 100 nL of AAV expressing GCaMP6m at a depth of 200 &#181;m at the center of each side of the SC.</li>
	<li>After ~3 weeks, implant a 5 mm diameter coverslip over the SC. Perform an oval craniotomy of 4 mm x 3 mm centered at the SC and thin the bone around it. Then, press the coverslip directly onto the SC and fix it with resin cement.</li>
	<li>Excite GCaMP6m with a blue light emitting diode (LED) with an excitation filter (469 nm &#177; 35 nm). Acquire images using a complementary metal oxide semiconductor (CMOS) camera after passing an emission filter (525 nm &#177; 39 nm) at 10 Hz. The camera covers an area of 5.36 mm x 2.85 mm at a spatial resolution of 2.63 &#181;m/pixel.</li>
	<li>Convolve each acquired image with a normalized box filter and downsample it to 1/4 of the original size. For each pixel in the image, define the response, as described in step 7.4. 
	NOTE: Provide postoperative care, as described in step 4.2.</li>
</ol>

Two-Photon and Wide-Field Calcium Imaging in the Superior Colliculus

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</ol>

The authors have nothing to disclose.

Critical steps in the protocol 
The most critical step is the craniotomy in steps 5.2 and 5.3. First, the bone at 0.5 mm posterior to the lambda is thick and has blood vessels inside, which can cause bleeding during the drilling process. Adequate gel foam should be prepared to stop the bleeding. Second, there is a good chance of angiorrhexis when removing the bone just above the transverse sinus. For troubleshooting, one alternative approach is to thin the bone inside the oval and remove it piece by piece. Another trick is to soak the bone prior to lifting to make the detachment of the dura from the bone easier and prevent bleeding.
Another critical step is the plug implantation in step 5.4, for which the whole process should last 1-2 min. For troubleshooting, one can first move the plug with the micromanipulator to an appropriate position in the ACSF and set the final position of the plug. Then, add the silicon adhesive and move the plug at an appropriate speed to the final position.
Significance 
This protocol has two merits. First, it provides details for imaging the posterior-medial SC at single-cell resolution with an intact cortex in wild-type mice, while in previous reports the overlaying cortex was aspirated to expose the anterior-lateral SC. Second, it details how to image the entire SC of partial-cortex mutant mice using the wide-field calcium imaging technique. These two approaches can be applied to image other brain regions in behaving animals at different scales.
Two alternative methods have been reported to image the posterior-medial SC without aspirating the overlaying cortex15,16. Schr&#246;der et al. had a circular 4 mm craniotomy and appled a stainless-steel tube with a washer and a glass coverslip at each end to expose the SC16. The larger craniotomy and more rigid material, compared to what we used, are more likely to cause bleeding during the implantation. In addition, because stainless steel is not biocompatible, it is more likely to cause infection for chronic imaging. Similarly, the glass coverslip used in Savier et al. was also more rigid and not biocompatible, and more likely cause bleeding and infections15. Another difference from these two methods is that our viral injection site is outside the imaging window, and thus the imaging quality is less affected by the potential damage at the injection site (Figure 5).
Limitations of the protocol 
This protocol provides details for imaging calcium responses in the superficial layer of the SC. Imaging deeper layers, for example using a prism26, is not covered. The two-photon calcium imaging approach is used for recording the neural activity in the posterior-medial SC and does not cover the anterior-lateral SC7,13. Entire SC calcium imaging with wide-field microscopy was applied to mutant partial-cortex mice. For imaging the entire SC of wide-type mice, one needs to aspirate the overlaying cortex. An alternative way to image the entire SC, without viral injection, is the intrinsic imaging technique; however, it has a lower signal-to-noise ratio6,8.

The superior colliculus (SC) is an important visual center in all vertebrates. In mammals, it receives direct inputs from the retina and the visual cortex1. While optical recording has been widely applied to the cortex2,3,4,5, its application in the SC is hindered by poor optical accesss6,7,8,9,10,11,12,13,14,15,16,17,18,19. The goal of this protocol is to provide details about two complementary methods for optical recording of the neural activity in the SC.
The SC is located beneath the cortex and transverse sinus, which limits optical access to the collicular neurons. One approach to overcome this limitation is to aspirate the overlaying cortex and expose the anterior-lateral SC7,9,10,13,14,19. However, because the SC receives cortical inputs, such an operation might affect how the SC neurons respond to visual stimuli. To overcome this limitation, we detail here an alternative protocol to image the superficial layer of the posterior-medial SC with a silicon plug, while leaving the cortex intact8,11. Specifically, to achieve single-cell resolution, we applied two-photon microscopy to image calcium responses in the posterior-medial SC of wild-type mice. In addition, to achieve broad coverage, we applied wide-field microscopy to image the entire SC of a mutant mouse whose posterior cortex has not developed20.
The two methods described in this protocol are complementary to each other. The two-photon calcium imaging without ablating the cortex is appropriate for recording neural activity at&#160;single-cell resolution with intact cortical inputs. The wide-field calcium imaging is appropriate for recording neural activity in the entire SC while sacrificing spatial resolution.

<table><tbody><tr><td>16x objective</td><td>Nikon</td><td></td><td></td></tr><tr><td>50-mm lens</td><td>Computar</td><td>M5018-MP2</td><td></td></tr><tr><td>5-mm coverslip</td><td>Warner instruments</td><td>CS-5R</td><td></td></tr><tr><td>bandpass filter</td><td>Chroma Technology</td><td>HQ575/250 m-2p</td><td></td></tr><tr><td>butyl cyanoacrylate</td><td>Vetbond, World Precision Instruments</td><td></td><td></td></tr><tr><td>camera for monitoring pupil</td><td>FLIR</td><td>BFS-U3-04S2M-CS</td><td></td></tr><tr><td>camera for widefield imaging</td><td>Basler</td><td>acA2000-165&amp;micro;m</td><td></td></tr><tr><td>corona treater</td><td>Electro-Technic Products</td><td>BD-20AC</td><td></td></tr><tr><td>dichroic</td><td>Chroma Technology</td><td>T600/200dcrb&amp;nbsp;</td><td></td></tr><tr><td>galvanometers</td><td>Cambridge Technology</td><td></td><td></td></tr><tr><td>glass bead sterilizer</td><td>RWD</td><td>RS1502</td><td></td></tr><tr><td>microdrill</td><td>RWD</td><td>78001</td><td></td></tr><tr><td>micromanipulator</td><td>Sutter Instruments</td><td>QUAD</td><td></td></tr><tr><td>photomultiplier tube</td><td>Hamamatsu</td><td>R3896</td><td></td></tr><tr><td>rotory encoder</td><td>USdigital</td><td>MA3-A10-125-N</td><td></td></tr><tr><td>self-curing dental adhesive resin cement&amp;nbsp;</td><td>SuperBond C&amp;amp;B, Sun Medical Co, Ltd. Moriyama, Japan</td><td></td><td></td></tr><tr><td>thermostatic heating pad&amp;nbsp;</td><td>RWD</td><td>69020</td><td></td></tr><tr><td>Ti:Sapphire laser</td><td>Spectra-Physics</td><td>Mai Tai HP DeepSee</td><td></td></tr><tr><td>translucent silicone adhesive&amp;nbsp;</td><td>Kwik-Sil, World Precision Instruments</td><td></td><td></td></tr><tr><td>treadmill</td><td>Xinglin Biology</td><td></td><td></td></tr><tr><td>&lt;strong&gt;Virus Strains&lt;/strong&gt;</td><td></td><td></td><td></td></tr><tr><td>rAAV2/9-hsyn-Gcamp6m</td><td>Vector Core at Chinese Institute for Brain Research, Beijing</td><td></td><td></td></tr><tr><td>&lt;strong&gt;Animals&lt;/strong&gt;</td><td></td><td></td><td></td></tr><tr><td>C57BL/6J wild type</td><td>Laboratory Animal Resource Center at Chinese Institute for Brain Research, Beijing</td><td></td><td></td></tr><tr><td>Emx1-Cre</td><td>The Jackson Laboratory&amp;nbsp;</td><td>5628</td><td></td></tr><tr><td>Pals1flox/wt</td><td>Christopher A. Walsh Lab</td><td></td><td></td></tr><tr><td>&lt;strong&gt;Software&lt;/strong&gt;</td><td></td><td></td><td></td></tr><tr><td>ImageJ</td><td>NIH Image</td><td></td><td></td></tr><tr><td>Labview</td><td>National Instruments</td><td></td><td></td></tr><tr><td>MATLAB</td><td>Mathworks</td><td></td><td></td></tr></tbody></table>

calcium imaging in mouse superior colliculus

The superior colliculus (SC), an evolutionarily conserved midbrain structure in all vertebrates, is the most sophisticated visual center before the emergence of the cerebral cortex. It receives direct inputs from ~30 types of retinal ganglion cells (RGCs), with each encoding a specific visual feature. It remains elusive whether the SC simply inherits retinal features or if additional and potentially de novo processing occurs in the SC. To reveal the neural coding of visual information in the SC, we provide here a detailed protocol to optically record visual responses with two complementary methods in awake mice. One method uses two-photon microscopy to image calcium activity at single-cell resolution without ablating the overlaying cortex, while the other uses wide-field microscopy to image the whole SC of a mutant mouse whose cortex is largely undeveloped. This protocol details these two methods, including animal preparation, viral injection, headplate implantation, plug implantation, data acquisition, and data analysis. The representative results show that the two-photon calcium imaging reveals visually evoked neuronal responses at single-cell resolution, and the wide-field calcium imaging reveals&#160;neural activity across the entire SC. By combining these two methods, one can reveal the neural coding in the SC at different scales, and such combination can also be applied to other brain regions.

Figures 1A,B show&#160;how to make the suction cup and the plugs, respectively. Figure 2 shows how to implant the plug successfully. After implanting the plug, the posterior-medial SC is exposed, as shown in Figure 2D. Figure 3 shows calcium responses of SC neurons from an example wild-type mouse imaged using two-photon microscopy. The triangular prism, which is easily captured under the microscope, can be used to locate the imaging site (Figure 3B). Visual responses are shown at&#160;single-cell resolution (Figures 3C ,Dand Video 1). Figure 4 shows calcium responses of SC neurons from an example partial-cortex mutant mouse imaged using wide-field microscopy. The acquired image was first downsampled to one-quarter of the original size (Figure 4B). Visually evoked calcium responses are shown at both sides of the SC (Figure 4C and Video 2). Figure 5 shows the expression of GCaMP6m in the SC at the injection site and the imaging site; there are less neurons at the injection site.
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/65181/65181fig01v4.jpg" /> 
Figure 1: Preparation. (A) Three steps for making the suction cup (steps 1.1-1.3); PBS inside the needle is not shown. (B) Three steps for making the plugs (steps 2.1-2.3). (C) Two types of headplate with an 8 mm diameter for wild-type mice. Two types of headplate with a 7 mm diameter for partial-cortex mutant mice. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/65181/65181fig01largev4.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/65181/65181fig02v2.jpg" /> 
Figure 2: Implantation of the plug (steps 5.2-5.5). (A) The bone is thinned. The surroundings are resin cement. (B) The bone and dura are removed to expose the inferior colliculus and cerebellum. (C) The plug is lowered, and it pushed the transverse sinus forward to expose the posterior-medial SC. The green dashed circle marks the coverslip. The blue dashed triangle marks the plug. The red dashed circle marks the suction cup. (D) The plug is fixed with butyl cyanoacrylate and resin cement. The image is overexposed to show the posterior-medial SC clearly. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/65181/65181fig02largev2.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 3" class="xfigimg" src="/files/ftp_upload/65181/65181fig03v4.jpg" /> 
Figure 3: Two-photon calcium imaging. (A) Schematic of the mouse brain anatomy after implanting the plug. The dark dashed line marks the outline of the SC. The yellow triangle indicates the triangular prism. The green color indicates the GCaMP expression. (B) A picture of the silicon triangular prism under the microscope before scanning. (C) An image of the fluorescence of GCamMP from SC neurons, measured by two-photon microscopy from an example wild-type mouse. (D) A standard deviation projection of the fluorescence across images (see Video 1). <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/65181/65181fig03largev4.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 4" class="xfigimg" src="/files/ftp_upload/65181/65181fig04v2.jpg" /> 
Figure 4: Wide-field calcium imaging. (A) A photo of the wide-field imaging setup. (B) An example raw image and the downsampled image. The dashed yellow line marks the outline of the SC. (C) A single frame and a standard deviation projection of the calcium responses of SC neurons, measured by wide-field microscopy from an example partial-cortex mouse (see Video 2). <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/65181/65181fig04largev2.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 5" class="xfigimg" src="/files/ftp_upload/65181/65181fig05v2.jpg" /> 
Figure 5: Coronal sections of a mouse brain at different distances from the injection site. The bottom row shows the high magnification of the boxes marked at the top row. The yellow dashed rectangle marks the injection site, which contains less neurons than the region on its right. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/65181/65181fig05largev2.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
Video 1: Raw and motion-corrected videos of the calcium responses of SC neurons to the visual stimulus in Video 3 by two-photon microscopy. The images were acquired at 4.8 Hz and displayed at 20 Hz. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/65181/two-photon-text.zip" target="_blank">Please click here to download this Video.</a>
Video 2: Raw and motion-corrected videos of the calcium responses of SC neurons to the visual stimulus in Video 3 by wide-field microscopy. The images were acquired at 10 Hz and displayed at 20 Hz. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/65181/wide field-text.zip" target="_blank">Please click here to download this Video.</a>
Video 3: A black full-screen bar (5&#176; wide) drifting in 12 different directions (50&#176;/s). <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/65181/MovingBars.zip" target="_blank">Please click here to download this Video.</a>

Watch this Scientific Journal Video about Unveiling Neural Coding and Mechanisms of Visual Processing in the Superior Colliculus at JoVE.com

Author Spotlight: Unveiling Neural Coding and Mechanisms of Visual Processing in the Superior Colliculus 

This protocol details the procedure for imaging calcium responses in the superior colliculus (SC) of awake mice, including imaging single-neuron activity with two-photon microscopy while leaving the cortex intact in wild-type mice, and imaging the entire SC with wide-field microscopy in partial-cortex mutant mice.

Calcium Imaging in Mouse Superior Colliculus

The superior colliculus (SC), an evolutionarily conserved midbrain structure in all vertebrates, is the most sophisticated visual center before the ...

calcium-imaging-in-mouse-superior-colliculus

Nanyang Technological University

Research

JoVE Journal

Neuroscience

3.3K Views.  Chinese Institute for Brain Research. Mijn onderzoek richt zich op neurale codering en mechanismen voor visuele verwerking. We proberen een antwoord te vinden op de vraag hoe verschillende informatie in de hersenen wordt gecodeerd, met name in de superieure colliculus onder de onderliggende neuromechanismen. Om visuele verwerking te begrijpen, combineren wetenschappers verschillende benaderingen, waaronder neuronale morfologie, antegrade en retrograde tracing, IL-sequencing, neuromodellering, machine learning, wordt goed begrepen...

Video: Auteur in de schijnwerpers: onthulling van neurale codering en mechanismen van visuele verwerking in de superieure colliculus 

Two-photon Calcium Imaging in Mice Navigating a Virtual Reality Environment

In recent years, two-photon imaging has become an invaluable tool in neuroscience, as it allows for chronic measurement of the activity of genetically identified cells during behavior1-6. Here we describe methods to perform two-photon imaging in mouse cortex while the animal navigates a virtual reality environment. We focus on the aspects of the experimental procedures that are key to imaging in a behaving animal in a brightly lit virtual environment. The key problems that arise in this experimental setup that we here address are: minimizing brain motion related artifacts, minimizing light leak from the virtual reality projection system, and minimizing laser induced tissue damage. We also provide sample software to control the virtual reality environment and to do pupil tracking. With these procedures and resources it should be possible to convert a conventional two-photon microscope for use in behaving mice.

Here we describe the experimental procedures involved in two-photon imaging of mouse cortex during behavior in a virtual reality environment.

In recent years, two-photon imaging has become an invaluable tool in neuroscience, as it allows for chronic measurement of the activity of genetically ...

Behavior

In Vivo 2-Photon Calcium Imaging in Layer 2/3 of Mice

To understand network dynamics of microcircuits in the neocortex, it is essential to simultaneously record the activity of a large number of neurons . In-vivo two-photon calcium imaging is the only method that allows one to record the activity of a dense neuronal population with single-cell resolution . The method consists in implanting a cranial imaging window, injecting a fluorescent calcium indicator dye that can be taken up by large numbers of neurons and finally recording the activity of neurons with time lapse calcium imaging using an in-vivo two photon microscope. Co-injection of astrocyte-specific dyes allows one to differentiate neurons from astrocytes. The technique can be performed in mice expressing fluorescent molecules in specific subpopulations of neurons to better understand the network interactions of different groups of cells.

To understand network dynamics of microcircuits in the neocortex, it is essential to simultaneously record the activity of a large number of neurons . In-vivo two-photon calcium imaging is the only method that allows one to record the activity of a dense neuronal population with single-cell resolution .

To understand network dynamics of microcircuits in the neocortex, it is essential to simultaneously record the activity of a large number of neurons .  ...

Biology

Imaging Calcium Responses in GFP-tagged Neurons of Hypothalamic Mouse Brain Slices

Despite an enormous increase in our knowledge about the mechanisms underlying the encoding of information in the brain, a central question concerning the precise molecular steps as well as the activity of specific neurons in multi-functional nuclei of brain areas such as the hypothalamus remain. This problem includes identification of the molecular components involved in the regulation of various neurohormone signal transduction cascades. Elevations of intracellular Ca2+ play an important role in regulating the sensitivity of neurons, both at the level of signal transduction and at synaptic sites.
New tools have emerged to help identify neurons in the myriad of brain neurons by expressing green fluorescent protein (GFP) under the control of a particular promoter. To monitor both spatially and temporally stimulus-induced Ca2+ responses in GFP-tagged neurons, a non-green fluorescent Ca2+ indicator dye needs to be used. In addition, confocal microscopy is a favorite method of imaging individual neurons in tissue slices due to its ability to visualize neurons in distinct planes of depth within the tissue and to limit out-of-focus fluorescence. The ratiometric Ca2+ indicator fura-2 has been used in combination with GFP-tagged neurons1. However, the dye is excited by ultraviolet (UV) light. The cost of the laser and the limited optical penetration depth of UV light hindered its use in many laboratories. Moreover, GFP fluorescence may interfere with the fura-2 signals2. Therefore, we decided to use a red fluorescent Ca2+ indicator dye. The huge Stokes shift of fura-red permits multicolor analysis of the red fluorescence in combination with GFP using a single excitation wavelength. We had previously good results using fura-red in combination with GFP-tagged olfactory neurons3. The protocols for olfactory tissue slices seemed to work equally well in hypothalamic neurons4. Fura-red based Ca2+ imaging was also successfully combined with GFP-tagged pancreatic &beta;-cells and GFP-tagged receptors expressed in HEK cells5,6. A little quirk of fura-red is that its fluorescence intensity at 650 nm decreases once the indicator binds calcium7. Therefore, the fluorescence of resting neurons with low Ca2+ concentration has relatively high intensity. It should be noted, that other red Ca2+-indicator dyes exist or are currently being developed, that might give better or improved results in different neurons and brain areas.

In this protocol, we update recent progress in imaging Ca2+ signals of GFP-tagged neurons in brain tissue slices using a red fluorescent Ca2+ indicator dye.

Despite an enormous increase in our knowledge about the mechanisms underlying the encoding of information in the brain, a central question concerning the ...

Immunohistochemical and Calcium Imaging Methods in Wholemount Rat Retina

In this paper we describe the tools, reagents, and the practical steps that are needed for: 1) successful preparation of wholemount retinas for immunohistochemistry and, 2) calcium imaging for the study of voltage gated calcium channel (VGCC) mediated calcium signaling in retinal ganglion cells. The calcium imaging method we describe circumvents issues concerning non-specific loading of displaced amacrine cells in the ganglion cell layer.

Immunohistochemistry protocols are used to study the localization of a specific protein in the retina. Calcium imaging techniques are employed to study calcium dynamics in retinal ganglion cells and their axons.

In this paper we describe the tools, reagents, and the practical steps that are needed for: 1) successful preparation of wholemount retinas for ...

Modification of a Colliculo-thalamocortical Mouse Brain Slice, Incorporating 3-D printing of Chamber Components and Multi-scale Optical Imaging

The ability of the brain to process sensory information relies on both ascending and descending sets of projections. Until recently, the only way to study these two systems and how they interact has been with the use of in vivo preparations. Major advances have been made with acute brain slices containing the thalamocortical and cortico-thalamic pathways in the somatosensory, visual, and auditory systems. With key refinements to our recent modification of the auditory thalamocortical slice1, we are able to more reliably capture the projections between most of the major auditory midbrain and forebrain structures: the inferior colliculus (IC), medial geniculate body (MGB), thalamic reticular nucleus (TRN), and the auditory cortex (AC). With portions of all these connections retained, we are able to answer detailed questions that complement the questions that can be answered with in vivo preparations. The use of flavoprotein autofluorescence imaging enables us to rapidly assess connectivity in any given slice and guide the ensuing experiment. Using this slice in conjunction with recording and imaging techniques, we are now better equipped to understand how information processing occurs at each point in the auditory forebrain as information ascends to the cortex, and the impact of descending cortical modulation. 3-D printing to build slice chamber components permits double-sided perfusion and broad access to networks within the slice and maintains the widespread connections key to fully utilizing this preparation.

Using multiple angles to cut the mouse pup brain, we improve upon a previously-described acute brain slice which captures the connections between most of the major auditory midbrain and forebrain structures.

The ability of the brain to process sensory information relies on both ascending and descending sets of projections. Until recently, the only way to study ...

Imaging Neural Activity in the Primary Somatosensory Cortex Using Thy1-GCaMP6s Transgenic Mice

The mammalian brain exhibits marked symmetry across the sagittal plane. However, detailed description of neural dynamics in symmetric brain regions in adult mammalian animals remains elusive. In this study, we describe an experimental procedure for measuring calcium dynamics through dual optical windows above bilateral primary somatosensory corticies (S1) in Thy1-GCaMP6s transgenic mice using 2-photon (2P) microscopy. This method enables recordings and quantifications of neural activity in bilateral mouse brain regions one at a time in the same experiment for a prolonged period in vivo. Key aspects of this method, which can be completed within an hour, include minimally invasive surgery procedures for creating dual optical windows, and the use of 2P imaging. Although we only demonstrate the technique in the S1 area, the method can be applied to other regions of the living brain facilitating the elucidation of structural and functional complexities of brain neural networks.

Imaging Neural Activity in the Primary Somatosensory Cortex Using Thy1-GCaMP6s Transgenic Mice

We describe an experimental procedure for measuring neuronal activity through dual optical windows above bilateral primary somatosensory corticies (S1) in Thy1-GCaMP6s transgenic mice using 2-photon (2P) microscopy in vivo.

The mammalian brain exhibits marked symmetry across the sagittal plane. However, detailed description of neural dynamics in symmetric brain regions in ...

In vivo Calcium Imaging of Mouse Geniculate Ganglion Neuron Responses to Taste Stimuli

Within the last ten years, advances in genetically encoded calcium indicators (GECIs) have promoted a revolution in in vivo functional imaging. Using calcium as a proxy for neuronal activity, these techniques provide a way to monitor the responses of individual cells within large neuronal ensembles to a variety of stimuli in real time. We, and others, have applied these techniques to image the responses of individual geniculate ganglion neurons to taste stimuli applied to the tongues of live anesthetized mice. The geniculate ganglion is comprised of the cell bodies of gustatory neurons innervating the anterior tongue and palate as well as some somatosensory neurons innervating the pinna of the ear. Imaging the taste-evoked responses of individual geniculate ganglion neurons with GCaMP has provided important information about the tuning profiles of these neurons in wild-type mice as well as a way to detect peripheral taste miswiring phenotypes in genetically manipulated mice. Here we demonstrate the surgical procedure to expose the geniculate ganglion, GCaMP fluorescence image acquisition, initial steps for data analysis, and troubleshooting. This technique can be used with transgenically encoded GCaMP, or with AAV-mediated GCaMP expression, and can be modified to image particular genetic subsets of interest (i.e., Cre-mediated GCaMP expression). Overall, in vivo calcium imaging of geniculate ganglion neurons is a powerful technique for monitoring the activity of peripheral gustatory neurons and provides complementary information to more traditional whole-nerve chorda tympani recordings or taste behavior assays.

Here we present how to expose the geniculate ganglion of a live, anesthetized laboratory mouse and how to use calcium imaging to measure the responses of ensembles of these neurons to taste stimuli, allowing for multiple trials with different stimulants. This allows for in depth comparisons of which neurons respond to which tastants.

Within the last ten years, advances in genetically encoded calcium indicators (GECIs) have promoted a revolution in in vivo functional imaging. Using ...

In vivo Calcium Imaging in Mouse Inferior Olive

Inferior olive (IO), a nucleus in the ventral medulla, is the only source of climbing fibers that form one of the two input pathways entering the cerebellum. IO has long been proposed to be crucial for motor control and its activity is currently considered to be at the center of many hypotheses of both motor and cognitive functions of the cerebellum. While its physiology and function have been relatively well studied on single-cell level in vitro, presently there are no reports on the organization of the IO network activity in living animals. This is largely due to the extremely challenging anatomical location of the IO, making it difficult to subject to conventional fluorescent imaging methods, where an optic path must be created through the entire brain located dorsally to the region of interest.
Here we describe an alternative method for obtaining state-of-the-art -level calcium imaging data from the IO network. The method takes advantage of the extreme ventral location of the IO and involves a surgical procedure for inserting a gradient-refractive index (GRIN) lens through the neck viscera to come into contact with the ventral surface of the calcium sensor GCaMP6s-expressing IO in anesthetized mice. A representative calcium imaging recording is shown to demonstrate the feasibility to record IO neuron activity after the surgery. While this is a non-survival surgery and the recordings must be conducted under anesthesia, it avoids damage to life-critical brainstem nuclei and allows conducting large variety of experiments investigating spatiotemporal activity patterns and input integration in the IO. This procedure with modifications could be used for recordings in other, adjacent regions of the ventral brainstem.

In vivo Calcium Imaging in Mouse Inferior Olive

We present a protocol to expose the brainstem of adult mouse from the ventral side. By using a gradient-refractive index lens with a miniature microscope, calcium imaging can be used to examine the activity of inferior olive neural somata in vivo.

Inferior olive (IO), a nucleus in the ventral medulla, is the only source of climbing fibers that form one of the two input pathways entering the ...

Brain Pericyte Calcium and Hemodynamic Imaging in Transgenic Mice In Vivo

Recent advances in protein biology and mouse genetics have made it possible to measure intracellular calcium fluctuations of brain cells in vivo and to correlate this with local hemodynamics. This protocol uses transgenic mice that have been prepared with a chronic cranial window and express the genetically encoded calcium indicator, RCaMP1.07, under the &#945;-smooth muscle actin promoter to specifically label mural cells, such as vascular smooth muscle cells and ensheathing pericytes. Steps are outlined on how to prepare a tail vein catheter for intravenous injection of fluorescent dyes to trace blood flow, as well as how to measure brain pericyte calcium and local blood vessel hemodynamics (diameter, red blood cell velocity, etc.) by two photon microscopy in vivo through the cranial window in ketamine/xylazine anesthetized mice. Finally, details are provided for the analysis of calcium fluctuations and blood flow movies via the image processing algorithms developed by Barrett et al. 2018, with an emphasis on how these processes can be adapted to other cellular imaging data.

Brain Pericyte Calcium and Hemodynamic Imaging in Transgenic Mice In Vivo

This protocol presents steps to acquire and analyze fluorescent calcium images from brain ensheathing pericytes and blood flow data from nearby blood vessels in anesthetized mice. These techniques are useful for studies of mural cell physiology and can be adapted to investigate calcium transients in any cell type.

Recent advances in protein biology and mouse genetics have made it possible to measure intracellular calcium fluctuations of brain cells in vivo and to ...

In Vivo Wide-Field and Two-Photon Calcium Imaging from a Mouse Using a Large Cranial Window

Wide-field calcium imaging from the mouse&#39;s neocortex allows one to observe cortex-wide neural activity related to various brain functions. On the other hand, two-photon imaging can resolve the activity of local neural circuits at the single-cell level. It is critical to make a large cranial window to perform multiple-scale analysis using both imaging techniques in the same mouse. To achieve this, one must remove a large section of the skull and cover the exposed cortical surface with transparent materials. Previously, glass skulls and polymer-based cranial windows have been developed for this purpose, but these materials are not easily fabricated. The present protocol describes a simple method for making a large cranial window consisting of commercially available polyvinylidene chloride (PVDC) wrapping film, a transparent silicone plug, and a cover glass. For imaging the dorsal surface of an entire hemisphere, the window size was approximately 6 x 3 mm2. Severe brain vibrations were not observed regardless of such a large window. Importantly, the condition of the brain surface did not deteriorate for more than one month. Wide-field imaging of a mouse expressing a genetically-encoded calcium indicator (GECI), GCaMP6f, specifically in astrocytes, revealed synchronized responses in a few millimeters. Two-photon imaging of the same mouse showed prominent calcium responses in individual astrocytes over several seconds. Furthermore, a thin layer of an adeno-associated virus was applied to the PVDC film and successfully expressed GECI in cortical neurons over the cranial window. This technique is reliable and cost-effective for making a large cranial window and facilitates the investigation of the neural and glial dynamics and their interactions during behavior at the macroscopic and microscopic levels.

In Vivo Wide-Field and Two-Photon Calcium Imaging from a Mouse Using a Large Cranial Window

The present protocol describes making a large (6 x 3 mm2) cranial window using food wrap, transparent silicone, and cover glass. This cranial window allows in vivo wide-field and two-photon calcium imaging experiments in the same mouse.

Wide-field calcium imaging from the mouse&#39;s neocortex allows one to observe cortex-wide neural activity related to various brain functions. On the ...

Calcium Imaging in Mouse Superior Colliculus

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