All animal procedures conformed to NIH guidelines and were approved by Cold Spring Harbor Laboratory Institutional Animal Care and Use Committee. Detailed methods are available as Supplementary material . To express GCaMP5 in corticalbulbar axons, adult mice were anesthetized with ketamine/xylazine and injected with a cocktail of AAV2.9 Synapsin-Cre and AAV2.9 EF1-DIO-GCaMP5 in the APC at least two weeks before imaging. To gain optical access to the OB, animals were anesthetized, the bone over the bulb was removed and replaced by a 3 mm coverslip (CS-3R, Warner Instruments). A titanium headbar was attached to the skull to fixate the animal during the imaging sessions. Mice were imaged using a Chameleon Ultra II Ti:Sapphire femtosecond pulsed laser (Coherent) coupled to a custom built multiphoton microscope.
Cs 3r
Manufactured by Warner Instruments
The CS-3R is a laboratory instrument manufactured by Warner Instruments. It serves as a chamber slide system for microscopic cell and tissue analysis. The device provides a controlled environment for specimens during observation and experimentation.
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Lab products found in correlation
8 protocols using cs 3r
1
In Vivo Imaging of Cortical Bulbar Axons
2
Hippocampal Cortical Imaging Window Implantation
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Cortical excavation and imaging window implantation were performed >10 days after the initial virus injection, according to published protocols10 (link),48 (link). A craniotomy (diameter 3 mm) was made centered at A/P −1.5 mm and M/L −1.5 mm for CA1/DG imaging and A/P −1.5 mm, M/L −2.5 mm for CA2/3 recordings. For implantations over CA2/3, the head of the mouse was tilted by 20°, so that the implantation plane was parallel to the lateral part of the pyramidal cell layer. Parts of the somatosensory cortex as well as posterior parietal association cortex were gently aspirated while irrigating with chilled saline. We continued aspiration until the external capsule was exposed. The outer part of the external capsule was then gently peeled away using fine forceps leaving the inner capsule and the hippocampus itself undamaged. The imaging window implant consisted of a 3 mm diameter coverslip (CS-3R, Warner Instruments) glued to the bottom of a stainless-steel cannula (3 mm diameter 1.2–1.5 mm height). The window was gradually lowered into the craniotomy using a forceps until the glass was in contact with the external capsule. The implant was then affixed to the skull using cyanoacrylate. Mice were allowed to recover from window implantation for 2-3 days.
3
Chronic Cranial Window Preparation
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After anesthetized with pentobarbital sodium, mice were maintained at a temperature of 37 °C under a heating pad throughout the cranial window surgery. A butterfly metal adaptor was first glued onto the skull, which was used for head fixation under 2PLSM. The cranial surgery was performed by drilling a 3-mm round window on the anterolateral parietal bone overlying the middle cerebral artery (MCA) territory. Afterward, the cranial window was sealed with a 3-mm round coverslip (Warner Instruments, CS-3R, Cat# 64-0720) with the instance adhesive (deli 502 super glue, Cat#No.7146). A small hole on the skull was drilled near the coverslip, when there is any drug need to be delivered. Two-photon imaging was performed on the second day after cranial window surgery. Otherwise, to avoid the influence of acute inflammation on vasomotion after surgery, in the cases of ischemic arteriolar vasomotion detection, the mice with a two-week chronic cranial window were checked and showed no difference between the mice with an acute cranial window.
4
Craniotomy and Chronic Imaging of Barrel Cortex
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Mice were anaesthetized with 2% isoflurane and placed into a stereotaxic frame. The body temperature of the mice was monitored and kept constant. Mice were kept on a thermal blanket, and their eyes were protected with artificial tear ointment. The skin above the skull was disinfected with povidone iodine. A craniotomy was made over the right barrel cortex (3 mm in diameter) using a biopsy punch and sealed with a glass coverslip (CS-3R Warner Instruments). A metal headplate (Luigs and Neuman) and the glass coverslip were then glued in place using dental cement (Superbond). Mice were allowed to recover for at least 2 weeks before doing habituation. Mice were habituated to the imaging setup by handling and training for 4-6 d.
5
Craniotomy and Coverslip Implantation
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After 2–3 weeks of viral injection surgeries, mice were anesthetized with 2% isoflurane in the same stereotaxic rig, and a titanium head plate was attached to the skull using dental cement. A 3 mm round craniotomy was made over the primary somatosensory cortex, and the dura was removed. The upper tangent of the 3 mm round glass coverslip (Warner Instruments, CS-3R) was placed at the same site as the viral injection and fixed to the skull using cyanoacrylate.
6
Imaging Window Implantation for In Vivo Cortical Recordings
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During a second surgery, taking place at least 7 days after the first one described in the previous section, an imaging window was implanted. A craniotomy (diameter 3 mm) was made at A/P −1.9 mm, M/L −1.8 mm. Under continuous irrigation with chilled saline, part of the somatosensory cortex and posterior parietal association cortex located above the hippocampus were progressively aspirated until the external capsule was exposed. The outer part of the external capsule was then gently peeled away using fine forceps, leaving the inner capsule and the hippocampus optically accessible, yet undamaged. The imaging window implant consisted of a 3-mm diameter coverslip (CS-3R, Warner Instruments) glued to the bottom of a stainless-steel cannula (3-mm diameter, 1.3-mm height). This window was gently lowered into the craniotomy using forceps until the coverslip was sitting on the external capsule. The implant was then fixed to the surrounding skull using cyanoacrylate. Mice were allowed to recover from window implantation for at least 4 days before any further experiment.
7
Implantation of Imaging Window for Cortical Imaging
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During a second surgery taking place at least 7 days after the first one (described in the previous section), an imaging window was implanted. Using the same anesthesia/analgesia protocol as described above, a craniotomy (diameter 3 mm) was drilled at A/P − 1.9 mm, M/L ± 1.8 mm. Under continuous irrigation with chilled saline, part of the somatosensory cortex and posterior parietal association cortex located above the hippocampus were progressively aspirated until the external capsule was exposed. The outer part of the external capsule was then gently peeled away using fine forceps, leaving the inner capsule and the hippocampus optically accessible, yet undamaged. The imaging window implant consisted of a 3-mm-wide coverslip (CS-3R, Warner Instruments) glued to the bottom of a stainless-steel cannula (3-mm diameter, 1.3-mm height). This window was gently lowered into the craniotomy using forceps until the coverslip was sitting on the external capsule. The implant was then fixed to the surrounding skull using cyanoacrylate. Mice were allowed to recover from window implantation for at least 4 days before any further experiment.
8
Customized Headpost and Cranial Window Implants
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Headpost implant design was adapted from Ghanbari et al. 55 , and consists of a custom-made Titanium or Stainless-steel head-plate, a 3D-printed frame, and three 0-80 screws to hold the frame to the head-plate.
We fabricated the head-plate with Titanium or Stainless-steel plate (McMaster-Carr) using a waterjet system (OMax), and 3D printed the frame using a ProJet MJP 2500 (3D Systems). Our design files can be found online (https://github.com/ckemere/TreadmillTracker/tree/master/UMinnHeadposts). We assembled the headpost implant after tapping the 3D printed frame with 0-80 tap and securing the headplate over the frame with three screws. The entire headpost is then stored in 70% Ethanol prior to surgery.
Cranial window fabrication procedure was adapted from Goldey et al. 56 . Windows were made of 2 stacked round coverslips (Warner Instruments # CS-3R, CS-4R, CS-5R) of different diameters. To fabricate the stacked windows, a 3 mm (or 4 mm) round coverslip was epoxied to a 4 mm (or 5 mm) cover slip using an optical adhesive (Norland Products Inc. e.g. # NOA 61, 71, 84) and cured using longwavelength UV light. To accommodate the large 5 mm stacked window, we cut off the right side of the 3D-printed frame to allow for extra space for the C&B Metabond to bind to the skull outside of the stacked cranial window. Fabricated stacked windows were stored in 70% ethanol prior to surgery.
We fabricated the head-plate with Titanium or Stainless-steel plate (McMaster-Carr) using a waterjet system (OMax), and 3D printed the frame using a ProJet MJP 2500 (3D Systems). Our design files can be found online (https://github.com/ckemere/TreadmillTracker/tree/master/UMinnHeadposts). We assembled the headpost implant after tapping the 3D printed frame with 0-80 tap and securing the headplate over the frame with three screws. The entire headpost is then stored in 70% Ethanol prior to surgery.
Cranial window fabrication procedure was adapted from Goldey et al. 56 . Windows were made of 2 stacked round coverslips (Warner Instruments # CS-3R, CS-4R, CS-5R) of different diameters. To fabricate the stacked windows, a 3 mm (or 4 mm) round coverslip was epoxied to a 4 mm (or 5 mm) cover slip using an optical adhesive (Norland Products Inc. e.g. # NOA 61, 71, 84) and cured using longwavelength UV light. To accommodate the large 5 mm stacked window, we cut off the right side of the 3D-printed frame to allow for extra space for the C&B Metabond to bind to the skull outside of the stacked cranial window. Fabricated stacked windows were stored in 70% ethanol prior to surgery.
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