This work was supported by the Ministry of Science and Technology, Taiwan (108-2320-B-002 -074, 109-2320-B-002-023-MY2).

All procedures performed in this study were approved by the National Taiwan University Animal Care and Use Committee (Approval No.: NTU-109-EL-00029 and NTU-108-EL-00158).
1. Assessment of the volume alteration of dental cement
NOTE: Changes in the volume of dental cement occur during the curing process. Test the volume changes of dental cement before implantation and baseplating. Researchers can test any brand of dental cement and use the brand with the lowest volume change to cement the baseplate. An example is shown in Figure 3.
<ol>
	<li>Weigh 0.5 g of each dental cement powder and mix it with its appropriate solution (1 mL). 
	NOTE: The recommended powder/liquid ratio in the dental cement instructions is 0.5 g of powder to 0.25 mL of the solution to obtain a solid form. This testing protocol dilutes the ratio to 0.5 g/mL so that the mixture can be aspirated in liquid form in order to measure the volume change in pipette tips.</li>
	<li>Use 0.1-10 &#181;L pipette tips to remove 2.5 &#181;L of the dental cement mixture, and then seal the pipette tip with a light curing glue.</li>
	<li>Place the tips on a rack, mark the top-most levels of the dental cement mixture, and measure the level of the dental cement mixture after 40 min (Supplementary video S1).</li>
	<li>In addition to monitoring the level changes during the dental cement curing in the tips, measure the remaining dry dental cement density. To calculate the density, measure the weight of the dental cement, and measure the volume with Archimedes&#39; principle.
	<ol>
		<li>In brief, place the remaining dry dental cement in a cup filled with water, and weigh the water that overflows.</li>
	</ol>
	</li>
	<li>Use the dental cement with the least shrinkage for all the protocols detailed below.</li>
</ol>
2. Anesthesia, surgical implantation, viral injection, and dummy baseplating
<ol>
	<li>Anesthesia 
	NOTE: The present report aims to optimize the surgery and baseplating procedure for the UCLA miniscope; therefore, it was assumed that the optimal viral titer for infecting the target brain regions was known. The methods for finding the optimal viral titer can be found in steps 1 to 5 of Resendez et al.8. Once the viral dilution has been optimized, surgical implantation can be initiated.
	<ol>
		<li>Place the mouse in an induction container and provide oxygen (100% at an airflow rate of 0.2 L/min). Preoxygenate the mouse for 5 to 10 min.</li>
		<li>Administer 5% isoflurane mixed with 100% oxygen (airflow rate: 0.2 L/min) until the mouse starts to lose its balance and is eventually anesthetized.</li>
		<li>Remove the mouse from the induction chamber and inject atropine (0.04 mg/kg; subcutaneous injection), to prevent the accumulation of saliva and buprenorphine (0.03 mg/kg; subcutaneous injection) as an analgesic.</li>
		<li>Shave the mouse&#39;s head.</li>
		<li>Wear a mask, sterile gown, and gloves. Prepare a sterile space and sterile surgical instruments for surgery. Stabilize the mouse&#39;s head (maintain anesthesia with 3 to 2.5% isoflurane during this step) on the stereotaxic apparatus. After the analgesic and anesthetic procedures, ensure that the ear bars securely stabilize the head. Provide thermal support.</li>
		<li>Ensure that the head is set in a straight position, sterilize the shaved head with betadine, and then spray the head with xylocaine (10%), a local analgesic.</li>
		<li>Apply veterinary ointment on the eyes to prevent dryness. 
		NOTE:&#160;Use ophthalmic ointment for eye protection during all anesthetic events to prevent ocular injury.</li>
		<li>Test the animal&#39;s deep pain reflex by pinching its hind paw. Once the animal shows no paw withdrawal reflex, proceed with surgical procedures.</li>
		<li>Make a small incision along the mid-sagittal plane of the skull (starting from approximately 2 mm anterior to the bregma and ending approximately 6 mm posterior to the bregma). Then, clean the connective tissue on the skull, and anchor three stainless steel screws onto the left frontal and interparietal bones.
		<ol>
			<li>At this point, reduce the isoflurane to 1-2%. Monitor the breathing rate during the whole surgical procedure. If the breathing rate is too slow (approximately 1/s, depending on the animal), decrease the concentration of isoflurane.</li>
		</ol>
		</li>
		<li>Execute a craniotomy for the relay lens implantation under a surgical microscope or stereomicroscope by using a burr drill with a 0.7 mm tip diameter. Use a micro-drill to grab the burr drill bit and draw an outline of the intended circle area (the full thickness of the calvarium does not need to be penetrated). 
		NOTE: The relay lens used in the present protocol had a diameter of 1.0 mm, a length ~ 9.0 mm a pitch of 1, and a working distance range of ~100 &#181;m - 300 &#181;m; therefore, the craniotomy had a diameter of 1.2 mm.
		<ol>
			<li>Gently deepen the outline until the dura is exposed.</li>
			<li>Prepare 3 mL of sterile saline in a 3 mL syringe and cool it in an ice bucket. Frequently rinse the exposed area with 0.1 mL of saline from the syringe to cool the area and prevent heat damage and hemorrhage.</li>
		</ol>
		</li>
		<li>Lightly pick and peel away the dura with a 27G needle.</li>
		<li>Put a marking on a 27 G blunt needle at 1 mm from the tip and use it to carefully aspirate the brain&#39;s cortex in order to create a window for lens implantation8,11,13,17 (Figure 2B1). If needed, make a blunt needle by grinding down the tip of a 27 G injection needle with sandpaper. Then, connect the needle to a syringe, connect the syringe to a tube, and connect the tube to a source of suction to create a vacuum. 
		NOTE: The relay lens is blunt and will compress the brain tissue when it is placed into the deep brain area. Thus, aspiration of the cortex reduces tissue damage due to relay lens implantation. The cortex is approximately 1 mm thick in mice but is difficult to visualize under a stereomicroscope. The mark creates a landmark to allow the depth of the needle to be determined more precisely than with a stereomicroscope alone. In addition, one can determine whether the cortex region has been reached or passed depending on the resistance; the top portion of the cortex feels relatively soft, while the subcortical region feels dense. Therefore, once the texture begins to feel denser, stop the aspiration (Figure 2B3).</li>
		<li>Prepare 3 mL of sterile saline in a 3 mL syringe and cool it in an ice bucket. Rinse the area with saline to stop the bleeding and decrease the possibility of brain edema.</li>
	</ol>
	</li>
	<li>Adeno-associated virus (AAV) injection 
	NOTE: AAV9-syn-jGCaMP7s-WPRE was used in the present experiment. jGCaMP7s is a genetically encoded calcium indicator that emits green fluorescence3. Because the present study used a wild-type mouse as a subject, a viral vector was needed to transfect the neurons with the green-fluorescent calcium indicator gene and enable the expression. Researchers using transgenic mice expressing the green-fluorescent calcium indicator as their subjects can skip protocol 2.2.
	<ol>
		<li>Mount the inner needle of a 20 G intravenous (i.v.) catheter onto the stereotaxic arm and slowly puncture (~100 - 200 &#181;m/min) the brain at the appropriate coordinates (the same coordinates used during relay lens implantation) (Figure 2B3).</li>
		<li>Lower the microinjection needle at a speed of 100-200 &#181;m/min into the ventral CA1 and infuse (~ 25 nL/min) 200 nL of the viral vector into the target region (-3.16 mm AP, 3.25 mm ML, and -3.50 mm DV from the bregma).</li>
		<li>Wait for 10 min to allow the virus to diffuse and to minimize the back flow.</li>
		<li>Withdraw (~100-200 &#181;m/min) the microinjection needle.</li>
	</ol>
	</li>
	<li>Relay lens implantation
	<ol>
		<li>Disinfect the relay lens with 75% alcohol10,16&#160;and then rinse with pyrogen-free saline. Soak the lens in cold pyrogen-free saline until implantation. 
		NOTE: A cold lens minimizes brain edema when placed into the brain.</li>
		<li>Hold the relay lens using a micro bulldog clamp (Figure 2B3) whose teeth have been covered with heat-shrink tubing. 
		NOTE: The bulldog clamp can firmly hold the lens without damaging it. Refer to Resendez et al.8 for the method to make the tubing-covered bulldog clamp.</li>
		<li>Slowly place (~100 - 200 &#181;m/min) the relay lens on top of the target region (-3.16 mm AP, 3.50 mm ML, and -3.50 mm DV from the bregma) and stabilize it with dental cement.</li>
	</ol>
	</li>
	<li>Dummy baseplating 
	NOTE: The purpose of &#34;dummy baseplating&#34; during the implantation surgery is to create a dental cement base in order to reduce the amount of dental cement that needs to be applied during the real baseplating procedure several weeks later. In this manner, the risk of a change in the volume of the dental cement is minimized.
	<ol>
		<li>Fasten the objective lens on the bottom of the miniscope and assemble the baseplate onto the miniscope (in this step, the assembly of the anchored objective lens, baseplate, and miniscope has not yet been placed over the mouse&#39;s skull).</li>
		<li>Wrap 10 cm of paraffin film around the outside of the baseplate (Figure 4A).</li>
		<li>Hold the miniscope with the stereotaxic arm probe using reusable adhesive clay.</li>
		<li>Align the objective lens on top of the relay lens with as little space between the lenses as possible (Figure 4B).</li>
		<li>Use dental cement to secure the positioning of the baseplate (such that the cement touches only the paraffin film).</li>
		<li>When the dental cement has dried, remove the film.</li>
		<li>Remove the baseplate and the miniscope. The dental cement base is hollow (Figure 4B).</li>
		<li>To protect the relay lens from daily activities and from being scratched by the mouse, seal the relay lens with some molding silicone rubber until the relay lens is covered, and cover the silicone rubber with a thin layer of dental cement (Figure 4B).</li>
		<li>Disengage the mouse from the stereotaxic instruments and place the mouse into a recovery chamber.</li>
		<li>Inject carprofen (5 mg/kg; subcutaneous injection) or meloxicam (1 mg/kg; subcutaneous injection) for analgesia and ceftazidime (25 mg/kg; subcutaneous injection) to prevent&#160;infection. 
		NOTE:&#160;The use of antibiotics is not an alternative to an aseptic technique. Perioperative antibiotics (i.e., ceftazidime) may be indicated in certain circumstances, such as long-duration surgeries or placement of chronic implants.</li>
		<li>Watch the animal until it regains sufficient consciousness to maintain sternal recumbency.</li>
		<li>House mice individually and inject meloxicam (1 mg/kg; subcutaneous injection; q24h) for one week to relieve postsurgical pain. Check the animal every day after surgery to make sure that it is eating, drinking, and defecating normally.</li>
		<li>Perform baseplating after 3 or more weeks.</li>
	</ol>
	</li>
</ol>
3. Baseplating
NOTE: Usually, the baseplating procedure (Figure 5) cannot be performed within 2-3 weeks of the initial procedure due to the recovery and viral incubation time. The induction procedures for baseplating are similar to those described in steps 2.1.1 to 2.1.8. However, baseplating is not an invasive procedure, and the animal requires only light sedation. Therefore, 0.8-1.2% isoflurane is sufficient, as long as the animal cannot move on the stereotaxic frame. In addition, relatively light isoflurane anesthesia can also help facilitate the expression of the fluorescence transients of neurons during monitoring8 (Supplementary video S2).
<ol>
	<li>Carefully cut the thin layer of the dental cement roof with a bone rongeur and remove the silicone rubber. Clean the surface of the relay lens with 75% alcohol.</li>
	<li>Fasten a set screw beside the baseplate to fix it to the bottom of the miniscope (Figure 5A1). 
	NOTE: The baseplate must be tightened securely under the bottom of the miniscope because the difference between a tightened and lightly fastened baseplate may cause an inconsistent distance.</li>
	<li>Adjust the focus slide to be approximately 2.7 - 3 mm from the main housing (Figure 5A2). 
	NOTE: Although the length can be further adjusted during the baseplating procedure, fix it to approximately 2.7 mm, because simultaneously adjusting the miniscope length and the optimal length between the implanted relay lens and preanchored objective lens is very confusing.</li>
	<li>Connect the miniscope to the data acquisition board and plug it into a USB 3.0 (or higher version) port on a computer.</li>
	<li>Run the data acquisition software developed by the UCLA team.</li>
	<li>Adjust the exposure to 255, the gain to 64x, and the excitation LED to 5%. These settings are recommended for initial baseplating; the specific settings will vary depending on the brain regions and the expression levels of the genetically encoded calcium indicator.</li>
	<li>Click the Connect button to connect the miniscope to the data acquisition software and watch the livestream.</li>
	<li>Align the miniscope objective lens to the relay lens by hand and begin searching for the fluorescence signals (Figure 5B1). 
	NOTE: Initially, searching for the fluorescence signal manually facilitates adjustments. Because an AAV system was used to express the genetically encoded calcium indicator, both the promoter type and AAV serotype affected the efficiency of transfection23,24. Researchers should need to check the expression of the calcium indicator by finding individual neurons that show changes in fluorescence before proceeding with future experiments.</li>
	<li>Drill the dental cement in any place where it blocks the miniscope (optional).</li>
	<li>Watch the live stream of the data acquisition software and use the margin of the relay lens as a landmark (Supplementary video S2). The margin of the relay lens appears as a moon-like gray circle on the monitor (Figure 5B1; white arrows).</li>
	<li>Once the relay lens is found, carefully adjust the various angles and distances of the miniscope until the fluorescence signals are found. 
	NOTE: The images of the neurons are whiter than the background. A precise neuronal shape indicates that the neurons lie perfectly on the focal plane of the relay lens. Round neurons suggest that they were near the focal plane. The neuronal shape is different across the brain, here ventral CA1 is shown as an example.</li>
	<li>If no individual neuron displays changes in fluorescence as a function of time, reseal the base with silicone rubber. Fluorescence is not observed because the incubation period is also dependent on the property of the virus23,24. Perform steps 3.1-3.12 after an additional week. (This is optional).</li>
	<li>Hold the miniscope at the optimal position and move the stereotaxic arm with a reusable adhesive clay towards the miniscope (Figure 5B2). Adhere the miniscope to the stereotaxic arm with reusable adhesive clay.</li>
	<li>Slightly adjust the x, y, and z arms to search for the best view (Supplementary video S2). Next, adjust the angle between the surface of the objective lens and relay lens by turning the ear bar, tooth bar, or reusable adhesive clay on the z-axis. Viral expression and lens implantation is successful when at least one cell displays fluorescence transients (Figure 6A; Supplementary video S2) during baseplating (which also depends on the area of the brain, such as the CA1). 
	NOTE: If the monitor shows only white cells but no transients, there is another cause (see Figure 6B,C, Supplementary videos S4,S5, the Results section, and the troubleshooting section).</li>
	<li>Cement the baseplate (Figure 5B2) with the dental cement.</li>
	<li>Monitor the region of interest during cementing to ensure that the optimal position does not change.</li>
	<li>Use the smallest possible amount of cement while still firmly gluing the baseplate onto the dental cement base (Figure 5B2). Apply a second and third layer of cement around the baseplate but be careful not to affect the GRIN lens. 
	NOTE: Applying cement to the baseplate is easier in this step since the base of the baseplate was formed during the dummy plating procedure.</li>
	<li>Carefully detach the miniscope from the baseplate when the cement is cured. Then, screw on a protective cap.</li>
	<li>Move the animal to a recovery cage. Watch the animal until it regains sufficient consciousness to maintain sternal recumbency.</li>
	<li>House mice individually. Make sure that it is eating, drinking, and defecating normally every day. Wait for 5 days for the mouse to fully recover and then check the calcium imaging. 
	NOTE: There is only a small amount of dental cement holding the baseplate. Therefore, if the baseplate has shifted considerably since the baseplating procedure, remove the dental cement with a drill perform the baseplating procedure again.</li>
	<li>Anesthetize (by intraperitoneal injection of a ketamine (87 mg/kg) /xylazine (13 mg/kg) mixture) and perfuse25 the animal when all experiments are done.</li>
</ol>

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<li>Gargiulo, S., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Mice+anesthesia%2C+analgesia%2C+and+care%2C+Part+I%3A+anesthetic+considerations+in+preclinical+research%5BTitle%5D)+AND+%22ILAR+journal+%2F+National+Research+Council%2C+Institute+of+Laboratory+Animal+Resources%22%5BJournal%5D)">Mice anesthesia, analgesia, and care, Part I: anesthetic considerations in preclinical research.</a> ILAR journal / National Research Council, Institute of Laboratory Animal Resources. 53 (1), 55-69 (2012).</li>
</ol>

This is not an industry-supported study. The authors report no financial conflicts of interest.

The present report describes a detailed experimental protocol for researchers using the two-lens UCLA Miniscope system. The tools designed in our protocol are relatively affordable for any laboratory that wishes to try in vivo calcium imaging. Some protocols, such as viral injection, lens implantation, dummy baseplating, and baseplating, could also be used for other versions of the miniscope system to improve the success rate of calcium imaging. Other than general problems with viral injection, and lens implanta...

Fluorescent activity reporters are ideal for imaging of the neuronal activity because they are sensitive and have large dynamic ranges1,2,3. Therefore, an increasing number of experiments are using fluorescence microscopy to directly observe neuronal activity1,2,3,4,5,6,7,8,9,10,11,12,13,14,15,16. The first miniaturized one-photon fluorescence microscope (miniscope) was designed in 2011 by Mark Schnitzer et al.5. This miniscope enables researchers to monitor the fluorescence dynamics of cerebellar cells in freely behaving animals5 (i.e., without any physical restraint, head restraint, sedation, or anesthesia to the animals). Currently, the technique can be applied to monitor superficial brain areas such as the cortex6,8,15,16; subcortical areas such as the dorsal hippocampus8,11,13,14 and striatum6,17; and deep brain areas such as the ventral hippocampus14, amygdala10,18, and hypothalamus8,12.
In recent years, several open-source miniscopes have been developed4,5,6,7,11,13,17,19. The miniscope can be economically assembled by researchers if they follow the step-by-step guidelines provided by the University of California, Los Angeles (UCLA) Miniscope team4,7,11,13. Because optical monitoring of neural activity is restricted by the limitations of light transmission7 to and from the neuronal population of interest, a miniscope was designed that requires an objective gradient refractive index (GRIN) lens (or objective lens) to be preanchored at the bottom of the miniscope to magnify the field of view that is relayed from a relay GRIN lens (or relay lens)6,7,8,10,16,17. This relay lens is implanted into the target brain region such that the fluorescence activity of the target brain region is relayed onto the surface of relay lens6,7,8,10,16,17. Approximately 1/4 of a full sinusoidal period of light travels through the objective GRIN lens (~ 0.25 pitch) (Figure 1A1), resulting in a magnified fluorescence image6,7. The objective lens is not always fixed at the bottom of the miniscope nor is the implantation of the relay lens necessary6,7,11,13,15. Specifically, there are two configurations: one with a fixed objective lens in the miniscope and a relay lens implanted in the brain8,10,12,14,16 (Figure 1B1) and another with just a removable objective lens6,7,11,13,15 (Figure 1B2). In the design based on the fixed objective and implanted relay lens combination, the fluorescence signals from the brain are brought to the top surface of the relay lens (Figure 1A1)7,8,10,12,14,16. Subsequently, the objective lens can magnify and transmit the visual field from the top surface of the relay lens (Figure 1A2). On the other hand, the removable objective GRIN lens design is more flexible, which means preimplantation of a relay lens into the brain is not mandatory (Figure 1B2)6,7,11,13,15. When using a miniscope based on a removable objective lens design, researchers still need to implant a lens into the target brain region but they can either implant an objective lens6,7,11,13,15 or a relay lens in the brain6,7. The choice of an objective or a relay lens for implantation determines the miniscope configuration that the researcher must use. For instance, the V3 UCLA Miniscope is based on a removable objective GRIN lens design. Researchers can choose either to directly implant an objective lens in the brain region of interest and mount the &#34;empty&#34; miniscope onto the objective lens6,7,11,13,15 (a one-lens system; Figure 1B2) or to implant a relay lens in the brain and mount a miniscope that is preanchored with an objective lens6,7 (a two-lens system; Figure 1B1). The miniscope then works as a fluorescence camera to capture livestream images of neuronal fluorescence produced by a genetically encoded calcium indicator1,2,3. After the miniscope is connected to a computer, these fluorescence images can be transferred to the computer and saved as video clips. Researchers can study neuronal activity by analyzing the relative changes in fluorescence with some analysis packages20,21 or write their codes for future analysis.
The V3 UCLA Miniscope provides flexibility for users to determine whether to image neuronal activity with a one- or two-lens system7. The choice of the recording system is based on the depth and size of the target brain area. In brief, a one-lens system can only image an area that is superficial (less than approximately 2.5 mm deep) and relatively large (larger than approximately 1.8 x 1.8 mm2) because the manufacturers only produce a certain size of objective lens. In contrast, a two-lens system can be applied to any target brain area. However, the dental cement for gluing the baseplate tends to cause misalignment with an altered distance between the objective and relay lenses, resulting in a poor image quality. If the two-lens system is being used, two working distances need to be precisely targeted to achieve the optimal imaging quality (Figure 1A). These two critical working distances are between the neurons and bottom surface of the relay lens, and between the top surface of the relay lens and the bottom surface of the objective lens (Figure 1A1). Any misalignment or misplacement of the lens outside of the working distance results in imaging failure (Figure 1C2). In contrast, the one-lens system requires only one precise working distance. However, objective lens size limits its application for monitoring of deep brain regions (the objective lens that fits the miniscope is approximately 1.8 ~ 2.0 mm6,11,13,15). Therefore, implantation of an objective lens is limited for the observation of the surface and relatively large brain regions, such as the cortex6,15 and dorsal cornu ammonis 1 (CA1) in mice11,13 . In addition, a large area of the cortex must be aspirated to target the dorsal CA111,13. Because of the limitation of the one-lens configuration that prevents imaging of deep brain regions, commercial miniscope systems offer only a combined objective lens/relay lens (two-lens) design. On the other hand, the V3 UCLA miniscope can be modified into either a one-lens or two-lens system because its objective lens is removable6,11,13,15. In other words, V3 UCLA miniscope users can take advantage of the removable lens by implanting it in the brain (creating a one-lens system), when performing experiments involving superficial brain observations (less than 2.5 mm in depth), or by preanchoring it in the miniscope and implanting a relay lens in the brain (creating a two-lens system), when performing experiments involving deep brain observations. The two-lens system can also be applied for superficially observing the brain, but the researcher must know the accurate working distances between the objective lens and relay lens. The main advantage of the one-lens system is that there is a decreased chance of missing the working distances than with a two-lens system, given that there are two working distances that need to be precisely targeted to achieve optimal imaging quality in the two-lens system (Figure 1A). Therefore, we recommend using a one-lens system for superficial brain observations. However, if the experiment requires imaging in the deep brain area, the researcher must learn to avoid misalignment of the two lenses.
The basic protocol for two-lens configuration of miniscopes for experiments includes lens implantation and baseplating8,10,16,17. Baseplating is the gluing of a baseplate onto an animal&#39;s head so that the miniscope can be eventually mounted on top of the animal and videotape the fluorescence signals of neurons (Figure 1B). This procedure involves using dental cement to glue the baseplate onto the skull (Figure 1C), but the shrinkage of dental cement can cause unacceptable changes in the distance between the implanted relay lens and the objective lens8,17. If the shifted distance between the two lenses is too large, cells cannot be brought into focus.
Detailed protocols for deep brain calcium imaging experiments using miniscopes have already been published8,10,16,17. The authors of these protocols have used the Inscopix system8,10,16 or other customized designs17 and have described the experimental procedures for viral selection, surgery, and baseplate attachment. However, their protocols cannot be precisely applied to other open-source systems, such as the V3 UCLA Miniscope system, NINscope6, and Finchscope19. Misalignment of the two lenses can occur during the recording in a two-lens configuration with a UCLA Miniscope due to the type of dental cement that is used to cement the baseplate to the skull8,17 (Figure 1C). The present protocol is needed because the distance between the implanted relay lens and the objective lens is prone to shift due to the undesirable shrinkage of dental cement during the baseplating procedure. During baseplating, the optimal working distance between the implanted relay lens and the objective lens must be found by adjusting the distance between the miniscope and the top of the relay lens, and the baseplate should then be glued at this ideal location. After the correct distance between the objective lens and the implanted relay lens is set, longitudinal measurements can be obtained at cellular resolution (Figure 1B; in vivo recording). Since the optimal range of working distances of a relay lens is small (50 - 350 &#181;m)4,8, excessive cement shrinkage during curing can make it difficult to keep the objective lens and the implanted relay lens within the appropriate range. The overall goal of this report is to provide a protocol to reduce the shrinkage problems8,17 that occur during the baseplating procedure and to increase the success rate of miniscope recordings of fluorescence signals in a two-lens configuration. Successful miniscope recording is defined as recording of a livestream of noticeable relative changes in the fluorescence of individual neurons in a freely behaving animal. Although different brands of dental cement have different shrinkage rates, researchers can select a brand that has been previously tested6,7,8,10,11,12,13,14,15,16,22. However, not every brand is easy to obtain in some countries/regions due to the import regulations for medical materials. Therefore, we have developed methods to test the shrinkage rates of available dental cements and, importantly provide an alternative protocol that minimizes the shrinkage problem. The advantage over the present baseplating protocol is an increase in the success rate of calcium imaging with tools and cement that can be easily obtained in laboratories. The UCLA miniscope is used as an example, but the protocol is also applicable to other miniscopes. In this report, we describe an optimized baseplating procedure and also recommend some strategies for fitting the UCLA miniscope two-lens system (Figure 2A). Both examples of successful implantation (n= 3 mice) and examples of failed implantation (n=2 mice) for the two-lens configuration with the UCLA miniscope are presented along with the discussions for the reasons of the successes and failures.

<table><tbody><tr><td>0.7-mm drill bit</td><td>&amp;nbsp;#19008-07</td><td>Fine Science Tools; USA</td><td>for surgery</td></tr><tr><td>0.1&amp;ndash;10 &amp;mu;l pipette tips</td><td>104-Q; QSP</td><td>Fisher Scientific; Singapore</td><td>for testing dental cement</td></tr><tr><td>20 G IV cathater</td><td>#SR-OX2032CA</td><td>Terumo Corporation; Tokyo, Japan</td><td>for surgery</td></tr><tr><td>27 G needle</td><td>AGANI, AN*2713R</td><td>Terumo Corporation; Tokyo, Japan</td><td>for surgery</td></tr><tr><td>AAV9-syn-jGCaMP7s-WPRE</td><td>#104487-AAV9; 1.5*10^13</td><td>Addgene viral prep; MA, USA</td><td>for viral injection</td></tr><tr><td>Atropine sulfate</td><td></td><td>Astart; Hsinchu, Taiwan</td><td>for surgery/dummy baseplating/baseplating</td></tr><tr><td>Baseplate</td><td>V3</td><td>http://miniscope.org</td><td>for dummy baseplating/baseplating</td></tr><tr><td>BLU TACK</td><td>#30840350</td><td>Bostik; Chelsea, Massachusetts, USA</td><td>Reusable adhesive clay; for surgery/dummy baseplating/baseplating</td></tr><tr><td>Bone Rongeur Friedman</td><td>13 cm</td><td>Diener; Tuttlingen, Germany</td><td>for baseplating</td></tr><tr><td>Buprenorphine</td><td></td><td>INDIVIOR; UK</td><td>for surgery</td></tr><tr><td>Carprofen</td><td>Rimadyl</td><td>Zoetis; Exton, PA</td><td>analgesia</td></tr><tr><td>Ceftazidime</td><td></td><td>Taiwan Biotech; Taiwan</td><td>prevent infection</td></tr><tr><td>Data Acquisition PCB for UCLA Miniscope</td><td>purchased on https://www.labmaker.org/collections/neuroscience/products/data-aquistion-system-daq</td><td></td><td>for baseplating</td></tr><tr><td>Dental cement set</td><td>Tempron</td><td>GC Corp; Tokyo, Japan</td><td>for testing dental cement</td></tr><tr><td>Dental cement set</td><td>Tokuso Curefast</td><td>Tokuyama Dental Corp.; Tokyo, Japan</td><td>for testing dental cement/surgery/dummy baseplating/baseplating</td></tr><tr><td>Dual Lab Standard with Mouse and Rat Adaptors</td><td>#51673</td><td>Stoelting Co; Illinois, USA</td><td>for surgery/dummy baseplating/baseplating</td></tr><tr><td>Duratear ointment</td><td></td><td>Alcon; Geneva, Switzerland</td><td>for surgery/dummy baseplating/baseplating</td></tr><tr><td>Ibuprofen</td><td></td><td>YungShin; Taiwan</td><td>analgesia</td></tr><tr><td>Isoflurane</td><td></td><td>Panion &amp;amp; BF Biotech INC.; Taoyuan, Taiwan</td><td>for surgery/dummy baseplating/baseplating</td></tr><tr><td>Inscopix</td><td>nVista System</td><td>Inscopix; Palo Alto, CA</td><td>for comparison with V3 UCLA Miniscope</td></tr><tr><td>Ketamine</td><td></td><td>Pfizer; NY, NY</td><td>for euthanasia</td></tr><tr><td>Normal saline</td><td></td><td></td><td>for surgery</td></tr><tr><td>Micro bulldog clamps</td><td>#12.102.04</td><td>Dimedo; Tuttlingen, Germany</td><td>for lens implantation</td></tr><tr><td>Microliter Microsyringes, 2.0 &amp;micro;L, 25 gauge</td><td>#88400</td><td>Hamilton; Bonaduz, Switzerland</td><td>for viral injection</td></tr><tr><td>Molding silicone rubber</td><td>ZA22 Thixo</td><td>Zhermack; Badia Polesine, Italy</td><td>for dummy baseplating</td></tr><tr><td>Objective Gradient index (GRIN) lens</td><td>#64519</td><td>Edmund Optics; NJ, USA</td><td>for dummy baseplating/baseplating</td></tr><tr><td>Parafilm</td><td>#PM996</td><td>Bemis; Neenah, USA</td><td>for dummy baseplating</td></tr><tr><td>Portable Suction</td><td>#DF-750</td><td>Doctor&#039;s Friend Medical Instrument Co., Inc., Taichung, Taiwan</td><td>for surgery</td></tr><tr><td>Relay GRIN lens</td><td>#1050-002177</td><td>Inscopix; Palo Alto, CA, USA</td><td>for dummy baseplating/baseplating</td></tr><tr><td>Stainless steel anchor screws</td><td>1.00 mm diameter, total length 3.00 mm</td><td></td><td>for surgery</td></tr><tr><td>Stereo microscope</td><td>#SL720</td><td>Sage Vison; New Taipei City, Taiwan</td><td>for surgery/dummy baseplating/baseplating</td></tr><tr><td>Stereotaxic apparatus</td><td>#51673</td><td>Stoelting; IL, USA</td><td>for surgery/dummy baseplating/baseplating</td></tr><tr><td>UV Cure Adhesive</td><td>#3321</td><td>Loctite; D&amp;uuml;sseldorf, Germany</td><td>for testing dental cement</td></tr><tr><td>V3 UCLA Miniscope</td><td>purchased on https://www.labmaker.org/products/miniscope-complete-set-of-components</td><td></td><td>for surgery/dummy baseplating/baseplating</td></tr><tr><td>Xylazine</td><td>X1126</td><td>Sigma-Aldrich; St. Louis, MO</td><td>for euthanasia</td></tr><tr><td>Xylocaine pump spray 10%</td><td></td><td>AstraZeneca; S&amp;ouml;dert&amp;auml;lje, Sweden</td><td>for surgery</td></tr></tbody></table>

using baseplating and a miniscope preanchored with an objective lens for calcium transient research in mice

Neuroscientists use miniature microscopes (miniscopes) to observe neuronal activity in freely behaving animals. The University of California, Los Angeles (UCLA) Miniscope team provides open resources for researchers to build miniscopes themselves. The V3 UCLA Miniscope is one of the most popular open-source miniscopes currently in use. It permits imaging of the fluorescence transients emitted from genetically modified neurons through an objective lens implanted on the superficial cortex (a one-lens system), or in deep brain areas through a combination of a relay lens implanted in the deep brain and an objective lens that is preanchored in the miniscope to observe the relayed image (a two-lens system). Even under optimal conditions (when neurons express fluorescence indicators and the relay lens has been properly implanted), a volume change of the dental cement between the baseplate and its attachment to the skull upon cement curing can cause misalignment with an altered distance between the objective and relay lenses, resulting in the poor image quality. A baseplate is a plate that helps mount the miniscope onto the skull and fixes the working distance between the objective and relay lenses. Thus, changes in the volume of the dental cement around the baseplate alter the distance between the lenses. The present protocol aims to minimize the misalignment problem caused by volume changes in the dental cement. The protocol reduces the misalignment by building an initial foundation of dental cement during relay lens implantation. The convalescence time after implantation is sufficient for the foundation of dental cement to cure the baseplate completely, so the baseplate can be cemented on this scaffold using as little new cement as possible. In the present article, we describe strategies for baseplating in mice to enable imaging of neuronal activity with an objective lens anchored in the miniscope.

Assessment of the dental cement volume alteration 
Since the volume of dental cement changes during the curing process, it may significantly impact the imaging quality, given that the working distance of a GRIN lens is approximately 50 to 350 µm4,8. Therefore, two commercially available dental cements were tested in this case, Tempron and Tokuso, before the implantation and baseplating procedure (Figure 5). Video...

Watch this Scientific Journal Video about Using Baseplating and a Miniscope Preanchored with an Objective Lens for Calcium Transient Research in Mice at JoVE.com

Using Baseplating and a Miniscope Preanchored with an Objective Lens for Calcium Transient Research in Mice

The shrinkage of dental cement during curing displaces the baseplate. This protocol minimizes the problem by creating an initial foundation of the dental cement that leaves space to cement the baseplate. Weeks later, the baseplate can be cemented in position on this scaffold using little new cement, therebyreducing shrinkage.

Neuroscientists use miniature microscopes (miniscopes) to observe neuronal activity in freely behaving animals. The University of California, Los Angeles ...

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Research

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Neuroscience

3.6K Views.  National Taiwan University. The shrinkage of dental cement during curing displaces the baseplate. This protocol minimizes the problem by creating an initial foundation of the dental cement that leaves space to cement the baseplate. Weeks later, the baseplate can be cemented in position on this scaffold using little new cement, therebyreducing shrinkage.

Video: Using Baseplating and a Miniscope Preanchored with an Objective Lens for Calcium Transient Research in Mice

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

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 ...

Stereotaxic Viral Injection and Gradient-Index Lens Implantation for Deep Brain In Vivo Calcium Imaging

A miniature fluorescence microscope (miniscope) is a potent tool for in vivo calcium imaging from freely behaving animals. It offers several advantages over conventional multi-photon calcium imaging systems: (1) compact; (2) light-weighted; (3) affordable; and (4) allows recording from freely behaving animals. This protocol describes brain surgeries for deep brain in vivo calcium imaging using a custom-developed miniscope recording system. The preparation procedure consists of three steps, including (1) stereotaxically injecting the virus at the desired brain region of a mouse brain to label a specific subgroup of neurons with genetically encoded calcium sensor; (2) implantation of gradient-index (GRIN) lens that can relay calcium image from deep brain region to the miniscope system; and (3) affixing the miniscope holder over the mouse skull where miniscope can be attached later. To perform in vivo calcium imaging, the miniscope is fastened onto the holder, and neuronal calcium images are collected along with simultaneous behavior recordings. The present surgery protocol is compatible with any commercial or custom-built single-photon and two-photon imaging systems for deep brain in vivo calcium imaging.

Stereotaxic Viral Injection and Gradient-Index Lens Implantation for Deep Brain In Vivo Calcium Imaging

Miniscope in vivo calcium imaging is a powerful technique to study neuronal dynamics and microcircuits in freely behaving mice. This protocol describes performing brain surgeries to achieve good in vivo calcium imaging using a miniscope.

A miniature fluorescence microscope (miniscope) is a potent tool for in vivo calcium imaging from freely behaving animals. It offers several advantages ...

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 ...

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

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.

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.

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

Real-Time Imaging for Neuron Activity in the Dentate Gyrus Cells of Mice 

In Vivo Calcium Imaging of Granule Cells in the Dentate Gyrus of Hippocampus in Mice

Real-time approaches are typically needed in studies of learning and memory, and in vivo calcium imaging provides the possibility to investigate neuronal activity in awake animals during behavior tasks. Since the hippocampus is closely associated with episodic and spatial memory, it has become an essential brain region in&#160;this field&#39;s research. In recent research, engram cells and place cells were studied by recording the neural activities in the hippocampal CA1 region using the miniature microscope in mice while performing behavioral tasks including open-field and linear track. Although the dentate gyrus is another important region in the hippocampus, it has rarely been studied with in vivo imaging due to its greater depth and difficulty for imaging. In this protocol, we present in detail a calcium imaging process, including how to inject the virus, implant a GRIN (Gradient-index) lens, and attach a base plate for imaging the dentate gyrus of the hippocampus. We further describe how to preprocess the calcium imaging data using MATLAB. Additionally, studies of other deep brain regions that require imaging may benefit from this method.

Author Spotlight: Investigating Neural Activity of Dentate Gyrus Granule Cells with Miniature Microscope

The dentate gyrus of the hippocampus carries out essential and distinct functions in learning and memory. This protocol describes a set of robust and efficient procedures for in vivo calcium imaging of granule cells in the dentate gyrus in freely moving mice.

Real-time approaches are typically needed in studies of learning and memory, and in vivo calcium imaging provides the possibility to investigate neuronal ...

Recording Neuronal Activity During Odor Plume Navigation with Miniscope Imaging

Miniscope Recording Calcium Signals at Hippocampus of Mice Navigating an Odor Plume

Mice navigate an odor plume with a complex spatiotemporal structure in the dark to find the source of odorants. This article describes a protocol to monitor behavior and record Ca2+ transients in dorsal CA1 stratum pyramidale neurons in the hippocampus (dCA1) in mice navigating an odor plume in a 50 cm x 50 cm x 25 cm odor arena. An epifluorescence miniscope focused through a gradient-index (GRIN) lens imaged Ca2+ transients in dCA1 neurons expressing the calcium sensor GCaMP6f in Thy1-GCaMP6f mice. The paper describes the behavioral protocol to train the mice to perform this odor plume navigation task in an automated odor arena. The methods include a step-by-step procedure for the surgery for GRIN lens implantation and baseplate placement for imaging GCaMP6f in CA1. The article provides information on real-time tracking of the mouse position to automate the start of the trials and delivery of a water reward. In addition, the protocol includes information on using an interface board to synchronize metadata describing the automation of the odor navigation task and frame times for the miniscope and a digital camera tracking mouse position. Moreover, the methods delineate the pipeline used to process GCaMP6f fluorescence movies by motion correction using the NorMCorre algorithm followed by identification of regions of interest with EXTRACT. Finally, the paper describes an artificial neural network approach to decode spatial paths from CA1 neural ensemble activity to predict mouse navigation of the odor plume.

Author Spotlight: Understanding Processing of Olfactory and Spatial Information by Brain with Real-Time Behavioral Analysis

This protocol investigates the brain-behavior relationship in hippocampal CA1 in mice navigating an odor plume. We provide a step-by-step protocol, including surgery to access imaging of the hippocampus, behavioral training, miniscope GCaMP6f recording and processing of the brain, and behavioral data to decode the mouse position from ROI neural activity.

Mice navigate an odor plume with a complex spatiotemporal structure in the dark to find the source of odorants. This article describes a protocol to ...

Visualization of Calcium Influx in Ventral Midbrain Neurons Derived from Mouse Embryos

Source:&#160;<a style='text-decoration:none;' href='https://app.jove.com/v/61481/quantifying-spontaneous-ca2-fluxes-and-their-downstream-effects-in-primary-mouse-midbrain-neurons'><span style='-webkit-text-decoration-skip:none;font-family:Arial,sans-serif;font-size:12pt;font-style:normal;font-variant:normal;font-weight:400;text-decoration-skip-ink:none;vertical-align:baseline;white-space:pre-wrap;'>Bancroft, E. A.,&#160;et al<span style='-webkit-text-decoration-skip:none;font-family:Arial,sans-serif;font-size:12pt;font-style:normal;font-variant:normal;font-weight:400;text-decoration-skip-ink:none;vertical-align:baseline;white-space:pre-wrap;'>. Quantifying Spontaneous Ca<span style='-webkit-text-decoration-skip:none;font-family:Arial,sans-serif;font-size:0.6em;font-style:normal;font-variant:normal;font-weight:400;text-decoration-skip-ink:none;vertical-align:baseline;white-space:pre-wrap;'>2+<span style='-webkit-text-decoration-skip:none;font-family:Arial,sans-serif;font-size:12pt;font-style:normal;font-variant:normal;font-weight:400;text-decoration-skip-ink:none;vertical-align:baseline;white-space:pre-wrap;'> Fluxes and their Downstream Effects in Primary Mouse Midbrain Neurons.&#160;J. Vis. Exp.<span style='-webkit-text-decoration-skip:none;font-family:Arial,sans-serif;font-size:12pt;font-style:normal;font-variant:normal;font-weight:400;text-decoration-skip-ink:none;vertical-align:baseline;white-space:pre-wrap;'> (2020)</a> &#160;This video demonstrates the real-time visualization of calcium influx in ventral midbrain neurons derived from mouse embryos. Using a viral vector to express a calcium indicator, the neurons are imaged under a confocal microscope to track changes in fluorescence intensity during spontaneous and neurotransmitter-induced calcium activity.

Source:&#160;Bancroft, E. A.,&#160;et al. Quantifying Spontaneous Ca2+ Fluxes and their Downstream Effects in Primary Mouse Midbrain Neurons.&#160;J. Vis. ...

Using Baseplating and a Miniscope Preanchored with an Objective Lens for Calcium Transient Research in Mice

Summary

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Visualization of Calcium Influx in Ventral Midbrain Neurons Derived from Mouse Embryos