http://miniscope.org/api.php?action=feedcontributions&user=Chrisl1026&feedformat=atomUCLA Miniscope - User contributions [en]2024-03-29T07:21:30ZUser contributionsMediaWiki 1.27.7http://miniscope.org/index.php?title=System_Assembly&diff=1763System Assembly2018-07-10T18:51:22Z<p>Chrisl1026: /* Soldering Coaxial Cable to CMOS Imaging Sensor PCB */</p>
<hr />
<div>This guide will take you through the assembly of the entire miniscope system.<br />
<br />
[[File:ScopeAssemblyParts.jpg|thumb|400px]]<br />
<br />
== Head Mounted Scope Assembly ==<br />
A detailed video is soon to come.<br />
<br />
#Examine '''Main Body''' under a microscope and remove any plastic burrs and obstructions to the light paths.<br />
#Press fit magnets into the 3 holes on the bottom of the '''Main Body''' making sure the polarity of the magnets match previously assembled scopes.<br />
#Slide the '''Achromatic Lens''' down the emission path until it sits flush against the aperture ring above the emission filter slot. The more curved of the two sides of the lens should face down toward the emission filter. Inspect fit under microscope and adjust if necessary.<br />
#With the coated surface facing the incoming light, use forceps to slide the ''Excitation Filter, Dichroic Mirror, and Emission Filter''' into their respective slots until their sides are flush with the Main Body. The black edges of the filters represent which edges should be blackened (optional).<br />
#Place the '''Half-Ball Lens''' in spherical opening (optical glue is optional). Inspect under a microscope to make sure the lens surface is flush with the plastic. Screw the Excitation LED PCB in place using 1mm self-tapping screws.<br />
#Screw the '''Filter Set Holder''' onto the '''Main Body''' using 2 to 3 1mm self-tapping screws.<br />
#Slide the '''Focusing Slider''' onto the '''Main Body'''. '''Make sure the two side holes have been tapped already with a 00-80 tap'''.<br />
#Epoxy, screw, or rubber band the '''CMOS Imaging Sensor PCB''' onto the '''Focusing Slider''' orienting the LED wires to the side of the scope with the '''Excitation LED PCB'''. <br />
<br />
[[File:ScopeAssembly2.png|center|900px]]<br />
<br clear=all><br />
<br />
{{#ev:youtube|https://youtu.be/p-RcVYbLlKc|640|center}}<br />
<br />
<br clear=all><br />
<br />
=== Filter Edge Blackening (NOT USUALLY NECESSARY) ===<br />
Our current version should not need this, but if you have light that leaks through the miniscope (i.e., you can see light being recorded even when the bottom of the miniscope is entirely covered), then blackening of the filters can fix this problem. We have noticed that if a scope has light leakage issues (excitation light making it to the CMOS imaging sensor) blackening the dichroic and emission filters' sides fixes the issue. <br />
<br />
Optical companies may be able to blacken the sides for you but it is also easy to do yourself. We have had the most success using Rustoleum Flat Black Enamel with a thin paintbrush.<br />
#Pour a few drops of the enamel into a plastic dish and let it sit out for a few minutes to thicken. <br />
#Holding the sides of the filter with forceps carefully apply a thin layer of the enamel to the 2 sides not in contact with the forceps. <br />
#Let dry for a few minutes before setting the filter down. <br />
#Wait a couple hours for the enamel to dry further then repeat step 2 on the other 2 sides of the filter.<br />
<br />
=== Soldering Coaxial Cable to CMOS Imaging Sensor PCB ===<br />
{{#ev:youtube|https://youtu.be/AZvfDosYdxc|640|center}}<br />
https://youtu.be/AZvfDosYdxc<br />
<br />
=== Soldering the solder-jumper on the bottom of the CMOS PCB (v3.2) ===<br />
If you are using version 3.2 of the CMOS imaging sensor PCB you will need to apply solder across the two half-circles on the bottom of the PCB.<br />
<br />
[[File:CMOS_Short_v3_2.png|center|600px]]<br />
<br />
=== Soldering SMA Connector to Coaxial Cable ===<br />
Below shows the easiest way to solder an SMA connector to the end of the coaxial cable. This can also be done using standard SMA wire connectors but it is more difficult and more prone to breaks. '''Make sure to test for shorts in the coax cable before plugging in and powering up the DAQ PCB. If a short is present, permanent damage can be done to the DAQ PCB.'''<br />
<br />
{{#ev:youtube|https://youtu.be/Jrn2oWERPpw|640|center}}<br />
<br />
=== Soldering LED to PCB ===<br />
{{#ev:youtube|8KWx6qv82ts|640|center}}<br />
<br />
'''Removing the silicone cover of the LED:''' try to slowly peal it off in 1 piece and make sure not to touch the white square (illumination surface) with forceps. It is fine to leave a small amount of silicone attached to the LED as long as it does not extend much higher than the actual surface of the LED.<br />
<br />
=== Soldering LED power lines ===<br />
[[File:LEDWire.png|thumb|300px]]<br />
{{#ev:youtube|https://youtu.be/AWg9-qTKF1Y|640|center}}<br />
<br />
'''Soldering the LED wires to the LED PCB:''' Once the LED PCB is attached to the scope body, the wires running off the LED PCB need to fit through a thin slot cut out of the scope body. This slot is only the width of the 2 solder pads on the LED PCB so care should be taken to solder the wires directly on top of the solder pads and have the wires extend straight off the PCB. It is helpful to use relatively thin wires, ideally with and outer diameter of 0.5mm or less.<br />
<br />
=== Baseplate Assembly ===<br />
#Inspect Baseplate for burrs.<br />
#Press fit the 3 magnets flush or slightly recessed into the Baseplate.<br />
#Tap the set screw hole with a 00-80 tap.<br />
<br />
[[File:BaseplateAssembly.png|center|400px]]<br />
<br />
== Data Acquisition System Assembly ==<br />
<br />
=== Note on version 3.x of the DAQ PCB ===<br />
If you have the newer version of the DAQ PCB (version 3.x) you can disregard the through-hole and coax cable connector soldering described below. The DAQ PCB v3.x comes fully soldered and just needs the EEPROM inserted and switch/jumper configuration set correctly. The coaxial cable from the head mounted scope will connect to the J4 (SMA connector closest to the center of the DAQ PCB) connector.<br />
<br />
<br />
[[File:DAQv3_x.png|center|500px]]<br />
<br />
=== Through-hole component assembly for version 2.x of PCB (USUALLY NOT NEEDED) ===<br />
If you decide to have the through-hole components assembled by an assembly house you can skip this section. Below is a picture highlighting the necessary through-hole components that need to be soldered in order for the DAQ PCB to function properly.<br />
<br />
[[File:DAQPCBThroughHole.png|center|500px]]<br />
<br />
*Description of components<br />
**SW4: Reset button the resets can reset the USB Host Controller<br />
**U5: EEPROM (memory that holds the DAQ firmware) socket. You can also solder the EEPROM IC directly to the board but I prefer using an IC socket so I can swap out the EEPROM if necessary<br />
**K1,2,3: Each are 2pin 0.1" headers<br />
**J9: A 3pin header used with a 2pin jumper to select power source for the microscope<br />
**J3,4,5: SMA connectors used for GPIO pins<br />
**J6: We currently solder a short coax cable with SMA connector to these pads. This will be updated soon to a replace this with a proper PCB footprint<br />
{{#ev:youtube|https://youtu.be/-BMkPaSKkk8|640|center}}<br />
<br />
=== Plugging in EEPROM ===<br />
Plug in the EEPROM into socket U5 making sure the orientation is correct (With the USB connector on the bottom the writing on the EEPROM should be right side up).<br />
<br />
[[File:EEPROM_Ori.png|center|500px]]<br />
<br />
=== Setting Jumpers ===<br />
Once all SMD and through-hole components are in place the switches and jumpers need to be properly set for uploading firmware and powering the microscope.<br />
<br />
Below shows the default configuration of the 3 SMD switches on the DAQ PCB.<br />
<br />
[[File:SwitchSettings.png|center|500px]]<br />
<br />
Below shows the possible K1, K2, and K3 jumper configurations.<br />
<br />
[[File:BootMode.png|center|500px]]<br />
<br />
The scope power jumper, J9, sets the power source powering the head mounted scope. In most cases the USB power configuration should be used and no DC power supply needs to be hooked up the the DC jack on the PCB.<br />
<br />
[[File:PowerJumper.png|center|500px]]<br />
<br />
=== Epoxy USB Connector ===<br />
If a USB cable is forced in or pulled out of the USB connector at an angle there is a chance the connector could get ripped off the PCB. While not necessary, we suggest adding epoxy between the USB connector housing and PCB to increase its mechanical stability and decrease the chances of damage. Care must be taken to avoid getting any epoxy inside the USB housing.<br />
<br />
== Cable Assembly ==<br />
The cabling between the head mounted scope and DAQ hardware is only a single coaxial cable. A coaxial, or coax, cable consists of an inner conducting wire surrounded by an insulating dielectric and then outer, generally grounded, shield. In our system the inner conductor carries power along with a data link and bidirectional control channel and the outer shield needs to be grounded. Our hardware dynamically adjusts for signal attenuation and small voltage drops across the cable but care should still be taken to minimize these loses. The videos above show how to solder the coax cable to the CMOS Imaging Sensor PCB and how to connectorize the other end with an SMA connector. Suggested cable can be found on our parts list but any coax cable with the properties listed below should work.<br />
<br />
Properties to look for in a coax cable are<br />
*50ohm impedance. This is absolutely necessary.<br />
*Light weight and highly flexible. We like to use coax cables with an outer diameter of 1.5mm or less. It is important to note that as the diameter of the cable decreases, so does the length it can support.<br />
*Handles bandwidths up to 1GHz. For short distances this requirement can be reduced.<br />
<br />
== What to do After System Assembly ==<br />
[[Software and Firmware Setup]]<br />
<br />
[[Initial Testing of Assembled Miniscopes]]</div>Chrisl1026http://miniscope.org/index.php?title=System_Assembly&diff=1762System Assembly2018-07-10T18:51:08Z<p>Chrisl1026: /* Soldering Coaxial Cable to CMOS Imaging Sensor PCB */</p>
<hr />
<div>This guide will take you through the assembly of the entire miniscope system.<br />
<br />
[[File:ScopeAssemblyParts.jpg|thumb|400px]]<br />
<br />
== Head Mounted Scope Assembly ==<br />
A detailed video is soon to come.<br />
<br />
#Examine '''Main Body''' under a microscope and remove any plastic burrs and obstructions to the light paths.<br />
#Press fit magnets into the 3 holes on the bottom of the '''Main Body''' making sure the polarity of the magnets match previously assembled scopes.<br />
#Slide the '''Achromatic Lens''' down the emission path until it sits flush against the aperture ring above the emission filter slot. The more curved of the two sides of the lens should face down toward the emission filter. Inspect fit under microscope and adjust if necessary.<br />
#With the coated surface facing the incoming light, use forceps to slide the ''Excitation Filter, Dichroic Mirror, and Emission Filter''' into their respective slots until their sides are flush with the Main Body. The black edges of the filters represent which edges should be blackened (optional).<br />
#Place the '''Half-Ball Lens''' in spherical opening (optical glue is optional). Inspect under a microscope to make sure the lens surface is flush with the plastic. Screw the Excitation LED PCB in place using 1mm self-tapping screws.<br />
#Screw the '''Filter Set Holder''' onto the '''Main Body''' using 2 to 3 1mm self-tapping screws.<br />
#Slide the '''Focusing Slider''' onto the '''Main Body'''. '''Make sure the two side holes have been tapped already with a 00-80 tap'''.<br />
#Epoxy, screw, or rubber band the '''CMOS Imaging Sensor PCB''' onto the '''Focusing Slider''' orienting the LED wires to the side of the scope with the '''Excitation LED PCB'''. <br />
<br />
[[File:ScopeAssembly2.png|center|900px]]<br />
<br clear=all><br />
<br />
{{#ev:youtube|https://youtu.be/p-RcVYbLlKc|640|center}}<br />
<br />
<br clear=all><br />
<br />
=== Filter Edge Blackening (NOT USUALLY NECESSARY) ===<br />
Our current version should not need this, but if you have light that leaks through the miniscope (i.e., you can see light being recorded even when the bottom of the miniscope is entirely covered), then blackening of the filters can fix this problem. We have noticed that if a scope has light leakage issues (excitation light making it to the CMOS imaging sensor) blackening the dichroic and emission filters' sides fixes the issue. <br />
<br />
Optical companies may be able to blacken the sides for you but it is also easy to do yourself. We have had the most success using Rustoleum Flat Black Enamel with a thin paintbrush.<br />
#Pour a few drops of the enamel into a plastic dish and let it sit out for a few minutes to thicken. <br />
#Holding the sides of the filter with forceps carefully apply a thin layer of the enamel to the 2 sides not in contact with the forceps. <br />
#Let dry for a few minutes before setting the filter down. <br />
#Wait a couple hours for the enamel to dry further then repeat step 2 on the other 2 sides of the filter.<br />
<br />
=== Soldering Coaxial Cable to CMOS Imaging Sensor PCB ===<br />
{{#ev:youtube|https://youtu.be/aXzErQn3U7g|640|center}}<br />
<br />
Updated:<br />
{{#ev:youtube|https://youtu.be/AZvfDosYdxc|640|center}}<br />
https://youtu.be/AZvfDosYdxc<br />
<br />
=== Soldering the solder-jumper on the bottom of the CMOS PCB (v3.2) ===<br />
If you are using version 3.2 of the CMOS imaging sensor PCB you will need to apply solder across the two half-circles on the bottom of the PCB.<br />
<br />
[[File:CMOS_Short_v3_2.png|center|600px]]<br />
<br />
=== Soldering SMA Connector to Coaxial Cable ===<br />
Below shows the easiest way to solder an SMA connector to the end of the coaxial cable. This can also be done using standard SMA wire connectors but it is more difficult and more prone to breaks. '''Make sure to test for shorts in the coax cable before plugging in and powering up the DAQ PCB. If a short is present, permanent damage can be done to the DAQ PCB.'''<br />
<br />
{{#ev:youtube|https://youtu.be/Jrn2oWERPpw|640|center}}<br />
<br />
=== Soldering LED to PCB ===<br />
{{#ev:youtube|8KWx6qv82ts|640|center}}<br />
<br />
'''Removing the silicone cover of the LED:''' try to slowly peal it off in 1 piece and make sure not to touch the white square (illumination surface) with forceps. It is fine to leave a small amount of silicone attached to the LED as long as it does not extend much higher than the actual surface of the LED.<br />
<br />
=== Soldering LED power lines ===<br />
[[File:LEDWire.png|thumb|300px]]<br />
{{#ev:youtube|https://youtu.be/AWg9-qTKF1Y|640|center}}<br />
<br />
'''Soldering the LED wires to the LED PCB:''' Once the LED PCB is attached to the scope body, the wires running off the LED PCB need to fit through a thin slot cut out of the scope body. This slot is only the width of the 2 solder pads on the LED PCB so care should be taken to solder the wires directly on top of the solder pads and have the wires extend straight off the PCB. It is helpful to use relatively thin wires, ideally with and outer diameter of 0.5mm or less.<br />
<br />
=== Baseplate Assembly ===<br />
#Inspect Baseplate for burrs.<br />
#Press fit the 3 magnets flush or slightly recessed into the Baseplate.<br />
#Tap the set screw hole with a 00-80 tap.<br />
<br />
[[File:BaseplateAssembly.png|center|400px]]<br />
<br />
== Data Acquisition System Assembly ==<br />
<br />
=== Note on version 3.x of the DAQ PCB ===<br />
If you have the newer version of the DAQ PCB (version 3.x) you can disregard the through-hole and coax cable connector soldering described below. The DAQ PCB v3.x comes fully soldered and just needs the EEPROM inserted and switch/jumper configuration set correctly. The coaxial cable from the head mounted scope will connect to the J4 (SMA connector closest to the center of the DAQ PCB) connector.<br />
<br />
<br />
[[File:DAQv3_x.png|center|500px]]<br />
<br />
=== Through-hole component assembly for version 2.x of PCB (USUALLY NOT NEEDED) ===<br />
If you decide to have the through-hole components assembled by an assembly house you can skip this section. Below is a picture highlighting the necessary through-hole components that need to be soldered in order for the DAQ PCB to function properly.<br />
<br />
[[File:DAQPCBThroughHole.png|center|500px]]<br />
<br />
*Description of components<br />
**SW4: Reset button the resets can reset the USB Host Controller<br />
**U5: EEPROM (memory that holds the DAQ firmware) socket. You can also solder the EEPROM IC directly to the board but I prefer using an IC socket so I can swap out the EEPROM if necessary<br />
**K1,2,3: Each are 2pin 0.1" headers<br />
**J9: A 3pin header used with a 2pin jumper to select power source for the microscope<br />
**J3,4,5: SMA connectors used for GPIO pins<br />
**J6: We currently solder a short coax cable with SMA connector to these pads. This will be updated soon to a replace this with a proper PCB footprint<br />
{{#ev:youtube|https://youtu.be/-BMkPaSKkk8|640|center}}<br />
<br />
=== Plugging in EEPROM ===<br />
Plug in the EEPROM into socket U5 making sure the orientation is correct (With the USB connector on the bottom the writing on the EEPROM should be right side up).<br />
<br />
[[File:EEPROM_Ori.png|center|500px]]<br />
<br />
=== Setting Jumpers ===<br />
Once all SMD and through-hole components are in place the switches and jumpers need to be properly set for uploading firmware and powering the microscope.<br />
<br />
Below shows the default configuration of the 3 SMD switches on the DAQ PCB.<br />
<br />
[[File:SwitchSettings.png|center|500px]]<br />
<br />
Below shows the possible K1, K2, and K3 jumper configurations.<br />
<br />
[[File:BootMode.png|center|500px]]<br />
<br />
The scope power jumper, J9, sets the power source powering the head mounted scope. In most cases the USB power configuration should be used and no DC power supply needs to be hooked up the the DC jack on the PCB.<br />
<br />
[[File:PowerJumper.png|center|500px]]<br />
<br />
=== Epoxy USB Connector ===<br />
If a USB cable is forced in or pulled out of the USB connector at an angle there is a chance the connector could get ripped off the PCB. While not necessary, we suggest adding epoxy between the USB connector housing and PCB to increase its mechanical stability and decrease the chances of damage. Care must be taken to avoid getting any epoxy inside the USB housing.<br />
<br />
== Cable Assembly ==<br />
The cabling between the head mounted scope and DAQ hardware is only a single coaxial cable. A coaxial, or coax, cable consists of an inner conducting wire surrounded by an insulating dielectric and then outer, generally grounded, shield. In our system the inner conductor carries power along with a data link and bidirectional control channel and the outer shield needs to be grounded. Our hardware dynamically adjusts for signal attenuation and small voltage drops across the cable but care should still be taken to minimize these loses. The videos above show how to solder the coax cable to the CMOS Imaging Sensor PCB and how to connectorize the other end with an SMA connector. Suggested cable can be found on our parts list but any coax cable with the properties listed below should work.<br />
<br />
Properties to look for in a coax cable are<br />
*50ohm impedance. This is absolutely necessary.<br />
*Light weight and highly flexible. We like to use coax cables with an outer diameter of 1.5mm or less. It is important to note that as the diameter of the cable decreases, so does the length it can support.<br />
*Handles bandwidths up to 1GHz. For short distances this requirement can be reduced.<br />
<br />
== What to do After System Assembly ==<br />
[[Software and Firmware Setup]]<br />
<br />
[[Initial Testing of Assembled Miniscopes]]</div>Chrisl1026http://miniscope.org/index.php?title=System_Assembly&diff=1761System Assembly2018-07-10T18:43:06Z<p>Chrisl1026: /* Soldering Coaxial Cable to CMOS Imaging Sensor PCB */</p>
<hr />
<div>This guide will take you through the assembly of the entire miniscope system.<br />
<br />
[[File:ScopeAssemblyParts.jpg|thumb|400px]]<br />
<br />
== Head Mounted Scope Assembly ==<br />
A detailed video is soon to come.<br />
<br />
#Examine '''Main Body''' under a microscope and remove any plastic burrs and obstructions to the light paths.<br />
#Press fit magnets into the 3 holes on the bottom of the '''Main Body''' making sure the polarity of the magnets match previously assembled scopes.<br />
#Slide the '''Achromatic Lens''' down the emission path until it sits flush against the aperture ring above the emission filter slot. The more curved of the two sides of the lens should face down toward the emission filter. Inspect fit under microscope and adjust if necessary.<br />
#With the coated surface facing the incoming light, use forceps to slide the ''Excitation Filter, Dichroic Mirror, and Emission Filter''' into their respective slots until their sides are flush with the Main Body. The black edges of the filters represent which edges should be blackened (optional).<br />
#Place the '''Half-Ball Lens''' in spherical opening (optical glue is optional). Inspect under a microscope to make sure the lens surface is flush with the plastic. Screw the Excitation LED PCB in place using 1mm self-tapping screws.<br />
#Screw the '''Filter Set Holder''' onto the '''Main Body''' using 2 to 3 1mm self-tapping screws.<br />
#Slide the '''Focusing Slider''' onto the '''Main Body'''. '''Make sure the two side holes have been tapped already with a 00-80 tap'''.<br />
#Epoxy, screw, or rubber band the '''CMOS Imaging Sensor PCB''' onto the '''Focusing Slider''' orienting the LED wires to the side of the scope with the '''Excitation LED PCB'''. <br />
<br />
[[File:ScopeAssembly2.png|center|900px]]<br />
<br clear=all><br />
<br />
{{#ev:youtube|https://youtu.be/p-RcVYbLlKc|640|center}}<br />
<br />
<br clear=all><br />
<br />
=== Filter Edge Blackening (NOT USUALLY NECESSARY) ===<br />
Our current version should not need this, but if you have light that leaks through the miniscope (i.e., you can see light being recorded even when the bottom of the miniscope is entirely covered), then blackening of the filters can fix this problem. We have noticed that if a scope has light leakage issues (excitation light making it to the CMOS imaging sensor) blackening the dichroic and emission filters' sides fixes the issue. <br />
<br />
Optical companies may be able to blacken the sides for you but it is also easy to do yourself. We have had the most success using Rustoleum Flat Black Enamel with a thin paintbrush.<br />
#Pour a few drops of the enamel into a plastic dish and let it sit out for a few minutes to thicken. <br />
#Holding the sides of the filter with forceps carefully apply a thin layer of the enamel to the 2 sides not in contact with the forceps. <br />
#Let dry for a few minutes before setting the filter down. <br />
#Wait a couple hours for the enamel to dry further then repeat step 2 on the other 2 sides of the filter.<br />
<br />
=== Soldering Coaxial Cable to CMOS Imaging Sensor PCB ===<br />
{{#ev:youtube|https://youtu.be/aXzErQn3U7g|640|center}}<br />
{{#ev:youtube|https://youtu.be/AZvfDosYdxc|640|center}}<br />
https://youtu.be/AZvfDosYdxc<br />
<br />
=== Soldering the solder-jumper on the bottom of the CMOS PCB (v3.2) ===<br />
If you are using version 3.2 of the CMOS imaging sensor PCB you will need to apply solder across the two half-circles on the bottom of the PCB.<br />
<br />
[[File:CMOS_Short_v3_2.png|center|600px]]<br />
<br />
=== Soldering SMA Connector to Coaxial Cable ===<br />
Below shows the easiest way to solder an SMA connector to the end of the coaxial cable. This can also be done using standard SMA wire connectors but it is more difficult and more prone to breaks. '''Make sure to test for shorts in the coax cable before plugging in and powering up the DAQ PCB. If a short is present, permanent damage can be done to the DAQ PCB.'''<br />
<br />
{{#ev:youtube|https://youtu.be/Jrn2oWERPpw|640|center}}<br />
<br />
=== Soldering LED to PCB ===<br />
{{#ev:youtube|8KWx6qv82ts|640|center}}<br />
<br />
'''Removing the silicone cover of the LED:''' try to slowly peal it off in 1 piece and make sure not to touch the white square (illumination surface) with forceps. It is fine to leave a small amount of silicone attached to the LED as long as it does not extend much higher than the actual surface of the LED.<br />
<br />
=== Soldering LED power lines ===<br />
[[File:LEDWire.png|thumb|300px]]<br />
{{#ev:youtube|https://youtu.be/AWg9-qTKF1Y|640|center}}<br />
<br />
'''Soldering the LED wires to the LED PCB:''' Once the LED PCB is attached to the scope body, the wires running off the LED PCB need to fit through a thin slot cut out of the scope body. This slot is only the width of the 2 solder pads on the LED PCB so care should be taken to solder the wires directly on top of the solder pads and have the wires extend straight off the PCB. It is helpful to use relatively thin wires, ideally with and outer diameter of 0.5mm or less.<br />
<br />
=== Baseplate Assembly ===<br />
#Inspect Baseplate for burrs.<br />
#Press fit the 3 magnets flush or slightly recessed into the Baseplate.<br />
#Tap the set screw hole with a 00-80 tap.<br />
<br />
[[File:BaseplateAssembly.png|center|400px]]<br />
<br />
== Data Acquisition System Assembly ==<br />
<br />
=== Note on version 3.x of the DAQ PCB ===<br />
If you have the newer version of the DAQ PCB (version 3.x) you can disregard the through-hole and coax cable connector soldering described below. The DAQ PCB v3.x comes fully soldered and just needs the EEPROM inserted and switch/jumper configuration set correctly. The coaxial cable from the head mounted scope will connect to the J4 (SMA connector closest to the center of the DAQ PCB) connector.<br />
<br />
<br />
[[File:DAQv3_x.png|center|500px]]<br />
<br />
=== Through-hole component assembly for version 2.x of PCB (USUALLY NOT NEEDED) ===<br />
If you decide to have the through-hole components assembled by an assembly house you can skip this section. Below is a picture highlighting the necessary through-hole components that need to be soldered in order for the DAQ PCB to function properly.<br />
<br />
[[File:DAQPCBThroughHole.png|center|500px]]<br />
<br />
*Description of components<br />
**SW4: Reset button the resets can reset the USB Host Controller<br />
**U5: EEPROM (memory that holds the DAQ firmware) socket. You can also solder the EEPROM IC directly to the board but I prefer using an IC socket so I can swap out the EEPROM if necessary<br />
**K1,2,3: Each are 2pin 0.1" headers<br />
**J9: A 3pin header used with a 2pin jumper to select power source for the microscope<br />
**J3,4,5: SMA connectors used for GPIO pins<br />
**J6: We currently solder a short coax cable with SMA connector to these pads. This will be updated soon to a replace this with a proper PCB footprint<br />
{{#ev:youtube|https://youtu.be/-BMkPaSKkk8|640|center}}<br />
<br />
=== Plugging in EEPROM ===<br />
Plug in the EEPROM into socket U5 making sure the orientation is correct (With the USB connector on the bottom the writing on the EEPROM should be right side up).<br />
<br />
[[File:EEPROM_Ori.png|center|500px]]<br />
<br />
=== Setting Jumpers ===<br />
Once all SMD and through-hole components are in place the switches and jumpers need to be properly set for uploading firmware and powering the microscope.<br />
<br />
Below shows the default configuration of the 3 SMD switches on the DAQ PCB.<br />
<br />
[[File:SwitchSettings.png|center|500px]]<br />
<br />
Below shows the possible K1, K2, and K3 jumper configurations.<br />
<br />
[[File:BootMode.png|center|500px]]<br />
<br />
The scope power jumper, J9, sets the power source powering the head mounted scope. In most cases the USB power configuration should be used and no DC power supply needs to be hooked up the the DC jack on the PCB.<br />
<br />
[[File:PowerJumper.png|center|500px]]<br />
<br />
=== Epoxy USB Connector ===<br />
If a USB cable is forced in or pulled out of the USB connector at an angle there is a chance the connector could get ripped off the PCB. While not necessary, we suggest adding epoxy between the USB connector housing and PCB to increase its mechanical stability and decrease the chances of damage. Care must be taken to avoid getting any epoxy inside the USB housing.<br />
<br />
== Cable Assembly ==<br />
The cabling between the head mounted scope and DAQ hardware is only a single coaxial cable. A coaxial, or coax, cable consists of an inner conducting wire surrounded by an insulating dielectric and then outer, generally grounded, shield. In our system the inner conductor carries power along with a data link and bidirectional control channel and the outer shield needs to be grounded. Our hardware dynamically adjusts for signal attenuation and small voltage drops across the cable but care should still be taken to minimize these loses. The videos above show how to solder the coax cable to the CMOS Imaging Sensor PCB and how to connectorize the other end with an SMA connector. Suggested cable can be found on our parts list but any coax cable with the properties listed below should work.<br />
<br />
Properties to look for in a coax cable are<br />
*50ohm impedance. This is absolutely necessary.<br />
*Light weight and highly flexible. We like to use coax cables with an outer diameter of 1.5mm or less. It is important to note that as the diameter of the cable decreases, so does the length it can support.<br />
*Handles bandwidths up to 1GHz. For short distances this requirement can be reduced.<br />
<br />
== What to do After System Assembly ==<br />
[[Software and Firmware Setup]]<br />
<br />
[[Initial Testing of Assembled Miniscopes]]</div>Chrisl1026http://miniscope.org/index.php?title=System_Assembly&diff=1555System Assembly2016-11-18T17:09:30Z<p>Chrisl1026: /* Head Mounted Scope Assembly */</p>
<hr />
<div>This guide will take you through the assembly of the entire miniscope system.<br />
<br />
[[File:ScopeAssemblyParts.jpg|thumb|400px]]<br />
<br />
== Head Mounted Scope Assembly ==<br />
A detailed video is soon to come.<br />
<br />
#Examine '''Main Body''' under a microscope and remove any plastic burrs and obstructions to the light paths.<br />
#Press fit magnets into the 3 holes on the bottom of the '''Main Body''' making sure the polarity of the magnets match previously assembled scopes.<br />
#Slide the '''Achromatic Lens''' down the emission path until it sits flush against the aperture ring above the emission filter slot. The more curved of the two sides of the lens should face down toward the emission filter. Inspect fit under microscope and adjust if necessary.<br />
#With the coated surface facing the incoming light, use forceps to slide the ''Excitation Filter, Dichroic Mirror, and Emission Filter''' into their respective slots until their sides are flush with the Main Body. The black edges of the filters represent which edges should be blackened (optional).<br />
#Place the '''Half-Ball Lens''' in spherical opening (optical glue is optional). Inspect under a microscope to make sure the lens surface is flush with the plastic. Screw the Excitation LED PCB in place using 1mm self-tapping screws.<br />
#Screw the '''Filter Set Holder''' onto the '''Main Body''' using 2 to 3 1mm self-tapping screws.<br />
#Slide the '''Focusing Slider''' onto the '''Main Body'''. '''Make sure the two side holes have been tapped already with a 00-80 tap'''.<br />
#Epoxy, screw, or rubber band the '''CMOS Imaging Sensor PCB''' onto the '''Focusing Slider''' orienting the LED wires to the side of the scope with the '''Excitation LED PCB'''. <br />
<br />
[[File:ScopeAssembly2.png|center|900px]]<br />
<br clear=all><br />
<br />
{{#ev:youtube|https://youtu.be/p-RcVYbLlKc|640|center}}<br />
<br />
<br clear=all><br />
<br />
=== Filter Edge Blackening (Suggested) ===<br />
While not strictly necessary, we suggest blackening the sides of the dichroic and emission filters. We have noticed that if a scope has light leakage issues (excitation light making it to the CMOS imaging sensor) blackening the dichroic and emission filters' sides fixes the issue. <br />
<br />
Optical companies may be able to blacken the sides for you but it is also easy to do yourself. We have had the most success using Rustoleum Flat Black Enamel with a thin paintbrush.<br />
#Pour a few drops of the enamel into a plastic dish and let it sit out for a few minutes to thicken. <br />
#Holding the sides of the filter with forceps carefully apply a thin layer of the enamel to the 2 sides not in contact with the forceps. <br />
#Let dry for a few minutes before setting the filter down. <br />
#Wait a couple hours for the enamel to dry further then repeat step 2 on the other 2 sides of the filter.<br />
<br />
=== Soldering Coaxial Cable to CMOS Imaging Sensor PCB ===<br />
{{#ev:youtube|https://youtu.be/aXzErQn3U7g|640|center}}<br />
<br />
=== Soldering the solder-jumper on the bottom of the CMOS PCB (v3.2) ===<br />
If you are using version 3.2 of the CMOS imaging sensor PCB you will need to apply solder across the two half-circles on the bottom of the PCB.<br />
<br />
[[File:CMOS_Short_v3_2.png|center|600px]]<br />
<br />
=== Soldering SMA Connector to Coaxial Cable ===<br />
Below shows the easiest way to solder an SMA connector to the end of the coaxial cable. This can also be done using standard SMA wire connectors but it more difficult and more prone to breaks.<br />
<br />
{{#ev:youtube|https://youtu.be/Jrn2oWERPpw|640|center}}<br />
<br />
=== Soldering LED to PCB ===<br />
{{#ev:youtube|8KWx6qv82ts|640|center}}<br />
<br />
'''Removing the silicone cover of the LED:''' try to slowly peal it off in 1 piece and make sure not to touch the white square (illumination surface) with forceps. It is fine to leave a small amount of silicone attached to the LED as long as it does not extend much higher than the actual surface of the LED.<br />
<br />
=== Soldering LED power lines ===<br />
[[File:LEDWire.png|thumb|300px]]<br />
{{#ev:youtube|https://youtu.be/AWg9-qTKF1Y|640|center}}<br />
<br />
'''Soldering the LED wires to the LED PCB:''' Once the LED PCB is attached to the scope body, the wires running off the LED PCB need to fit through a thin slot cut out of the scope body. This slot is only the width of the 2 solder pads on the LED PCB so care should be taken to solder the wires directly on top of the solder pads and have the wires extend straight off the PCB. It is helpful to use relatively thin wires, ideally with and outer diameter of 0.5mm or less.<br />
<br />
=== Baseplate Assembly ===<br />
#Inspect Baseplate for burrs.<br />
#Press fit the 3 magnets flush or slightly recessed into the Baseplate.<br />
#Tap the set screw hole with a 00-80 tap.<br />
<br />
[[File:BaseplateAssembly.png|center|400px]]<br />
<br />
== Data Acquisition System Assembly ==<br />
We generally have all surface mount (SMD) components assembled on the DAQ PCB by a third party PCB assembly house leaving only the through-hole components to be assembled in lab. It is possible to have the assembly house place both SMD and through-hole components but it is more expensive and through-hole components are relatively easy to solder. A good through-hole soldering tutorial can be found [https://learn.sparkfun.com/tutorials/how-to-solder---through-hole-soldering here].<br />
<br />
=== Through-hole component assembly ===<br />
If you decide to have the through-hole components assembled by an assembly house you can skip this section. Below is a picture highlighting the necessary through-hole components that need to be soldered in order for the DAQ PCB to function properly.<br />
<br />
[[File:DAQPCBThroughHole.png|center|500px]]<br />
<br />
*Description of components<br />
**SW4: Reset button the resets can reset the USB Host Controller<br />
**U5: EEPROM (memory that holds the DAQ firmware) socket. You can also solder the EEPROM IC directly to the board but I prefer using an IC socket so I can swap out the EEPROM if necessary<br />
**K1,2,3: Each are 2pin 0.1" headers<br />
**J9: A 3pin header used with a 2pin jumper to select power source for the microscope<br />
**J3,4,5: SMA connectors used for GPIO pins<br />
**J6: We currently solder a short coax cable with SMA connector to these pads. This will be updated soon to a replace this with a proper PCB footprint<br />
{{#ev:youtube|https://youtu.be/-BMkPaSKkk8|640|center}}<br />
<br />
=== Plugging in EEPROM ===<br />
Plug in the EEPROM into socket U5 making sure the orientation is correct (With the USB connector on the bottom the writing on the EEPROM should be right side up).<br />
<br />
[[File:EEPROM_Ori.png|center|500px]]<br />
<br />
=== Setting Jumpers ===<br />
Once all SMD and through-hole components are in place the switches and jumpers need to be properly set for uploading firmware and powering the microscope.<br />
<br />
Below shows the default configuration of the 3 SMD switches on the DAQ PCB.<br />
<br />
[[File:SwitchSettings.png|center|500px]]<br />
<br />
Below shows the possible K1, K2, and K3 jumper configurations.<br />
<br />
[[File:BootMode.png|center|500px]]<br />
<br />
The scope power jumper, J9, sets the power source powering the head mounted scope. In most cases the USB power configuration should be used and no DC power supply needs to be hooked up the the DC jack on the PCB.<br />
<br />
[[File:PowerJumper.png|center|500px]]<br />
<br />
=== Epoxy USB Connector ===<br />
If a USB cable is forced in or pulled out of the USB connector at an angle there is a chance the connector could get ripped off the PCB. While not necessary, we suggest adding epoxy between the USB connector housing and PCB to increase its mechanical stability and decrease the chances of damage. Care must be taken to avoid getting any epoxy inside the USB housing.<br />
<br />
== Cable Assembly ==<br />
The cabling between the head mounted scope and DAQ hardware is only a single coaxial cable. A coaxial, or coax, cable consists of an inner conducting wire surrounded by an insulating dielectric and then outer, generally grounded, shield. In our system the inner conductor carries power along with a data link and bidirectional control channel and the outer shield needs to be grounded. Our hardware dynamically adjusts for signal attenuation and small voltage drops across the cable but care should still be taken to minimize these loses. The videos above show how to solder the coax cable to the CMOS Imaging Sensor PCB and how to connectorize the other end with an SMA connector. Suggested cable can be found on our parts list but any coax cable with the properties listed below should work.<br />
<br />
Properties to look for in a coax cable are<br />
*50ohm impedance. This is absolutely necessary.<br />
*Light weight and highly flexible. We like to use coax cables with an outer diameter of 1.5mm or less. It is important to note that as the diameter of the cable decreases, so does the length it can support.<br />
*Handles bandwidths up to 1GHz. For short distances this requirement can be reduced.<br />
<br />
== What to do After System Assembly ==<br />
[[Software and Firmware Setup]]<br />
<br />
[[Initial Testing of Assembled Miniscopes]]</div>Chrisl1026http://miniscope.org/index.php?title=Animal_Behavior_Guide&diff=1554Animal Behavior Guide2016-11-16T01:17:13Z<p>Chrisl1026: /* Setting imaging focal plane & LED intensity */</p>
<hr />
<div>Here are some tips to for using the miniscopes to run experiments in freely moving mice.<br />
<br />
<br />
== Experimental timeline ==<br />
Plan you experiments schedule accordingly. Here is a typical timeline for getting mice ready for experiments in freely moving mice.<br />
<br />
:Week 1: virus injection surgery<br />
:Week 2: GRIN lens implant surgery<br />
:Week 3: recovery<br />
:Week 4: base plate surgery + acclimation to experimenters/handling<br />
:Week 5: habituation to wearing miniscopes & setting imaging focal plane<br />
:Week 6: begin experiment!<br />
<br />
== Acclimation to experimenters/Handling ==<br />
We typically wait at least a day after base plate surgery to begin handling. We recommend that mice become well acclimated to the experimenters prior to mounting miniscopes on awake moving mice. It is important that the mice feel comfortable with the experimenters as stress during the mounting of the miniscope during the experiment can cause adverse effects for the mice and thus, experiment. After gently handling the animals a few times, acclimate the mice to getting something placed on and off their heads by gently taking the protective caps on and off their base plate (that is cemented on the mice's heads). The experimenters should be able to quickly remove the cap without much struggle from the animal before continuing onto the habituation phase.<br />
<br />
<br clear=all><br />
<br />
{{#ev:youtube|https://youtu.be/RjH8wx-_ebM|640|center}}<br />
<br />
<br clear=all><br />
<br />
{{#ev:youtube|https://youtu.be/WRIWRfkt5q8|640|center}}<br />
<br />
<br clear=all><br />
<br />
== Habituation to wearing miniscopes ==<br />
Once the mice (and experimenters) are comfortable with the cap removal, habituate the mice to wearing the miniscopes while freely moving. To habituate, one experimenter should calmly hold the mouse and remove the cap, while the second experimenter replaces the cap with the miniscope. The miniscope should easily snap into place with the magnets. The second experimenter should gently hold the sides of the base plate with one hand while securing the miniscope with the set screw with the other hand. Especially during the first time, the animal may struggle. If the animal is struggling too much, let go and calm the animal. If the animal resists and struggles TOO much while you are holding onto the base plate, it can lead to the dental cement holding the base plate to come off the skull. When the animal is calm, try to hold the base plate and insert the set screw to secure the miniscope again. Once the miniscope is secured, let the animal explore your habituation environment of choice. We typically just let them move freely around in their home cage (even if there are other mice in the cage). Depending on your experimental demands, you will need to habituate to different levels of movement. If the experiment only requires mice to walk around and explore, we recommend habituating for at least 3 days for 10 minutes each day. If the experiment requires the mice to run swiftly (e.g. down a linear track), then we recommend habituating for at least 5 days for 10-30 minutes each day.<br />
<br />
<br clear=all><br />
<br />
{{#ev:youtube|https://youtu.be/39ZNCih7ooE|640|center}}<br />
<br />
<br clear=all><br />
<br />
{{#ev:youtube|https://youtu.be/JRg1eaPF6Ao|640|center}}<br />
<br />
<br clear=all><br />
<br />
== Setting imaging focal plane & LED intensity ==<br />
We recommend setting the focal plane of imaging and LED intensity prior to the experimental day. This can be done during the habituation sessions. Move the focus slider along the neck of the body of the miniscope until you've reached your optimal focal plane of the brain. To set the focal plane, tighten the set screw into the focus slider until the focus slider is secure. Be careful of over-tightening, as the screw can make indents into to the plastic material of the neck of the body. Once the focal plane is set, do not change it for the duration of your experiment if you need to image the same set of cells across sessions. Typically, we have found that for some mice, they can use the same miniscope set at a certain focal plane. For other mice, they need the miniscope to be set to a different height.<br />
<br />
During this session, you may also want to find the optimal level of LED intensity for imaging. Make sure that the there is enough blue light to see both the firing of the cells as well as some blood vessels to be used as landmarks to align frames. We recommend taking a video during the habituation session with your settings and analyzing the video to make sure the settings are optimal prior to the start of the experiment. Also, take a snapshot of the brain that you will use to reference for all future recording sessions. That way, you can begin your experiment with confidence and ease!<br />
<br />
<br clear=all><br />
<br />
{{#ev:youtube|https://youtu.be/k1MbzqaRqK8|640|center}}<br />
<br />
<br clear=all><br />
<br />
== Starting an experiment ==<br />
Here is an example checklist that can be used.<br />
<br />
*Check if there is enough space on the hard drive for the data files that will be collected throughout the experiment. If you are using a behavioral camera, don't forget to include that as well.<br />
*Check and clean the imaging sensor before screwing it on the focus slider. If there is dust or dirt, use lens paper and alcohol to gently clean the sensor. This typically only needs to be done once prior to securing onto the focus slide. If the imaging sensor is being removed between sessions, make sure to screw it back onto the focus slide in the same orientation!<br />
*Connect the miniscope cable with the DAQ, connect the USB 3.0 cable with the DAQ and computer, and open the Miniscope Control application on the computer.<br />
*Make sure the 3 lights on the DAQ turn on and the red light on the miniscope turns on.<br />
*Make sure that the miniscope is flush against the base plate or as much as possible and the miniscope is not tilted and the set screw is not in the way.<br />
*Compare the live image you see with the snapshot of the same region you took during the habituation. Try to match it up as best as possible. The blood vessels, contrast, brightness should all look similar. If it doesn't then troubleshoot as to why it is different.<br />
*Gently tighten the set screw to secure the miniscope onto the base plate. Stop once the screw makes contact with the plastic of the miniscope. Over tightening will make an indentation into the plastic body and cause damage. To check if the miniscope is secure, GENTLY wiggle and tug on the miniscope while attached to the base plate while watching the imaging of the the brain. Moving the miniscope should not cause movement of the image on the video.<br />
*Make sure to have enough slack on the cable so the mice can move freely but not so much that the mice can chew on the cable. If you're not using a commutator, attend to the possible twisting of the cables while the animals are moving.<br />
*Check to see that the write speed of the computer exceeds the FPS collected. <br />
*After the trial, visually inspect that data folder to make sure the data was correctly collected.<br />
*Back up your data!!!</div>Chrisl1026http://miniscope.org/index.php?title=Animal_Behavior_Guide&diff=1553Animal Behavior Guide2016-11-16T01:14:53Z<p>Chrisl1026: /* Habituation to wearing miniscopes */</p>
<hr />
<div>Here are some tips to for using the miniscopes to run experiments in freely moving mice.<br />
<br />
<br />
== Experimental timeline ==<br />
Plan you experiments schedule accordingly. Here is a typical timeline for getting mice ready for experiments in freely moving mice.<br />
<br />
:Week 1: virus injection surgery<br />
:Week 2: GRIN lens implant surgery<br />
:Week 3: recovery<br />
:Week 4: base plate surgery + acclimation to experimenters/handling<br />
:Week 5: habituation to wearing miniscopes & setting imaging focal plane<br />
:Week 6: begin experiment!<br />
<br />
== Acclimation to experimenters/Handling ==<br />
We typically wait at least a day after base plate surgery to begin handling. We recommend that mice become well acclimated to the experimenters prior to mounting miniscopes on awake moving mice. It is important that the mice feel comfortable with the experimenters as stress during the mounting of the miniscope during the experiment can cause adverse effects for the mice and thus, experiment. After gently handling the animals a few times, acclimate the mice to getting something placed on and off their heads by gently taking the protective caps on and off their base plate (that is cemented on the mice's heads). The experimenters should be able to quickly remove the cap without much struggle from the animal before continuing onto the habituation phase.<br />
<br />
<br clear=all><br />
<br />
{{#ev:youtube|https://youtu.be/RjH8wx-_ebM|640|center}}<br />
<br />
<br clear=all><br />
<br />
{{#ev:youtube|https://youtu.be/WRIWRfkt5q8|640|center}}<br />
<br />
<br clear=all><br />
<br />
== Habituation to wearing miniscopes ==<br />
Once the mice (and experimenters) are comfortable with the cap removal, habituate the mice to wearing the miniscopes while freely moving. To habituate, one experimenter should calmly hold the mouse and remove the cap, while the second experimenter replaces the cap with the miniscope. The miniscope should easily snap into place with the magnets. The second experimenter should gently hold the sides of the base plate with one hand while securing the miniscope with the set screw with the other hand. Especially during the first time, the animal may struggle. If the animal is struggling too much, let go and calm the animal. If the animal resists and struggles TOO much while you are holding onto the base plate, it can lead to the dental cement holding the base plate to come off the skull. When the animal is calm, try to hold the base plate and insert the set screw to secure the miniscope again. Once the miniscope is secured, let the animal explore your habituation environment of choice. We typically just let them move freely around in their home cage (even if there are other mice in the cage). Depending on your experimental demands, you will need to habituate to different levels of movement. If the experiment only requires mice to walk around and explore, we recommend habituating for at least 3 days for 10 minutes each day. If the experiment requires the mice to run swiftly (e.g. down a linear track), then we recommend habituating for at least 5 days for 10-30 minutes each day.<br />
<br />
<br clear=all><br />
<br />
{{#ev:youtube|https://youtu.be/39ZNCih7ooE|640|center}}<br />
<br />
<br clear=all><br />
<br />
{{#ev:youtube|https://youtu.be/JRg1eaPF6Ao|640|center}}<br />
<br />
<br clear=all><br />
<br />
== Setting imaging focal plane & LED intensity ==<br />
We recommend setting the focal plane of imaging and LED intensity prior to the experimental day. This can be done during the habituation sessions. Move the focus slider along the neck of the body of the miniscope until you've reached your optimal focal plane of the brain. To set the focal plane, tighten the set screw into the focus slider until the focus slider is secure. Be careful of over-tightening, as the screw can make indents into to the plastic material of the neck of the body. Once the focal plane is set, do not change it for the duration of your experiment if you need to image the same set of cells across sessions. Typically, we have found that for some mice, they can use the same miniscope set at a certain focal plane. For other mice, they need the miniscope to be set to a different height.<br />
<br />
During this session, you may also want to find the optimal level of LED intensity for imaging. Make sure that the there is enough blue light to see both the firing of the cells as well as some blood vessels to be used as landmarks to align frames. We recommend taking a video during the habituation session with your settings and analyzing the video to make sure the settings are optimal prior to the start of the experiment. Also, take a snapshot of the brain that you will use to reference for all future recording sessions. That way, you can begin your experiment with confidence and ease!<br />
<br />
== Starting an experiment ==<br />
Here is an example checklist that can be used.<br />
<br />
*Check if there is enough space on the hard drive for the data files that will be collected throughout the experiment. If you are using a behavioral camera, don't forget to include that as well.<br />
*Check and clean the imaging sensor before screwing it on the focus slider. If there is dust or dirt, use lens paper and alcohol to gently clean the sensor. This typically only needs to be done once prior to securing onto the focus slide. If the imaging sensor is being removed between sessions, make sure to screw it back onto the focus slide in the same orientation!<br />
*Connect the miniscope cable with the DAQ, connect the USB 3.0 cable with the DAQ and computer, and open the Miniscope Control application on the computer.<br />
*Make sure the 3 lights on the DAQ turn on and the red light on the miniscope turns on.<br />
*Make sure that the miniscope is flush against the base plate or as much as possible and the miniscope is not tilted and the set screw is not in the way.<br />
*Compare the live image you see with the snapshot of the same region you took during the habituation. Try to match it up as best as possible. The blood vessels, contrast, brightness should all look similar. If it doesn't then troubleshoot as to why it is different.<br />
*Gently tighten the set screw to secure the miniscope onto the base plate. Stop once the screw makes contact with the plastic of the miniscope. Over tightening will make an indentation into the plastic body and cause damage. To check if the miniscope is secure, GENTLY wiggle and tug on the miniscope while attached to the base plate while watching the imaging of the the brain. Moving the miniscope should not cause movement of the image on the video.<br />
*Make sure to have enough slack on the cable so the mice can move freely but not so much that the mice can chew on the cable. If you're not using a commutator, attend to the possible twisting of the cables while the animals are moving.<br />
*Check to see that the write speed of the computer exceeds the FPS collected. <br />
*After the trial, visually inspect that data folder to make sure the data was correctly collected.<br />
*Back up your data!!!</div>Chrisl1026http://miniscope.org/index.php?title=Animal_Behavior_Guide&diff=1552Animal Behavior Guide2016-11-16T01:12:41Z<p>Chrisl1026: /* Acclimation to experimenters/Handling */</p>
<hr />
<div>Here are some tips to for using the miniscopes to run experiments in freely moving mice.<br />
<br />
<br />
== Experimental timeline ==<br />
Plan you experiments schedule accordingly. Here is a typical timeline for getting mice ready for experiments in freely moving mice.<br />
<br />
:Week 1: virus injection surgery<br />
:Week 2: GRIN lens implant surgery<br />
:Week 3: recovery<br />
:Week 4: base plate surgery + acclimation to experimenters/handling<br />
:Week 5: habituation to wearing miniscopes & setting imaging focal plane<br />
:Week 6: begin experiment!<br />
<br />
== Acclimation to experimenters/Handling ==<br />
We typically wait at least a day after base plate surgery to begin handling. We recommend that mice become well acclimated to the experimenters prior to mounting miniscopes on awake moving mice. It is important that the mice feel comfortable with the experimenters as stress during the mounting of the miniscope during the experiment can cause adverse effects for the mice and thus, experiment. After gently handling the animals a few times, acclimate the mice to getting something placed on and off their heads by gently taking the protective caps on and off their base plate (that is cemented on the mice's heads). The experimenters should be able to quickly remove the cap without much struggle from the animal before continuing onto the habituation phase.<br />
<br />
<br clear=all><br />
<br />
{{#ev:youtube|https://youtu.be/RjH8wx-_ebM|640|center}}<br />
<br />
<br clear=all><br />
<br />
{{#ev:youtube|https://youtu.be/WRIWRfkt5q8|640|center}}<br />
<br />
<br clear=all><br />
<br />
== Habituation to wearing miniscopes ==<br />
Once the mice (and experimenters) are comfortable with the cap removal, habituate the mice to wearing the miniscopes while freely moving. To habituate, one experimenter should calmly hold the mouse and remove the cap, while the second experimenter replaces the cap with the miniscope. The miniscope should easily snap into place with the magnets. The second experimenter should gently hold the sides of the base plate with one hand while securing the miniscope with the set screw with the other hand. Especially during the first time, the animal may struggle. If the animal is struggling too much, let go and calm the animal. If the animal resists and struggles TOO much while you are holding onto the base plate, it can lead to the dental cement holding the base plate to come off the skull. When the animal is calm, try to hold the base plate and insert the set screw to secure the miniscope again. Once the miniscope is secured, let the animal explore your habituation environment of choice. We typically just let them move freely around in their home cage (even if there are other mice in the cage). Depending on your experimental demands, you will need to habituate to different levels of movement. If the experiment only requires mice to walk around and explore, we recommend habituating for at least 3 days for 10 minutes each day. If the experiment requires the mice to run swiftly (e.g. down a linear track), then we recommend habituating for at least 5 days for 10-30 minutes each day.<br />
<br />
== Setting imaging focal plane & LED intensity ==<br />
We recommend setting the focal plane of imaging and LED intensity prior to the experimental day. This can be done during the habituation sessions. Move the focus slider along the neck of the body of the miniscope until you've reached your optimal focal plane of the brain. To set the focal plane, tighten the set screw into the focus slider until the focus slider is secure. Be careful of over-tightening, as the screw can make indents into to the plastic material of the neck of the body. Once the focal plane is set, do not change it for the duration of your experiment if you need to image the same set of cells across sessions. Typically, we have found that for some mice, they can use the same miniscope set at a certain focal plane. For other mice, they need the miniscope to be set to a different height.<br />
<br />
During this session, you may also want to find the optimal level of LED intensity for imaging. Make sure that the there is enough blue light to see both the firing of the cells as well as some blood vessels to be used as landmarks to align frames. We recommend taking a video during the habituation session with your settings and analyzing the video to make sure the settings are optimal prior to the start of the experiment. Also, take a snapshot of the brain that you will use to reference for all future recording sessions. That way, you can begin your experiment with confidence and ease!<br />
<br />
== Starting an experiment ==<br />
Here is an example checklist that can be used.<br />
<br />
*Check if there is enough space on the hard drive for the data files that will be collected throughout the experiment. If you are using a behavioral camera, don't forget to include that as well.<br />
*Check and clean the imaging sensor before screwing it on the focus slider. If there is dust or dirt, use lens paper and alcohol to gently clean the sensor. This typically only needs to be done once prior to securing onto the focus slide. If the imaging sensor is being removed between sessions, make sure to screw it back onto the focus slide in the same orientation!<br />
*Connect the miniscope cable with the DAQ, connect the USB 3.0 cable with the DAQ and computer, and open the Miniscope Control application on the computer.<br />
*Make sure the 3 lights on the DAQ turn on and the red light on the miniscope turns on.<br />
*Make sure that the miniscope is flush against the base plate or as much as possible and the miniscope is not tilted and the set screw is not in the way.<br />
*Compare the live image you see with the snapshot of the same region you took during the habituation. Try to match it up as best as possible. The blood vessels, contrast, brightness should all look similar. If it doesn't then troubleshoot as to why it is different.<br />
*Gently tighten the set screw to secure the miniscope onto the base plate. Stop once the screw makes contact with the plastic of the miniscope. Over tightening will make an indentation into the plastic body and cause damage. To check if the miniscope is secure, GENTLY wiggle and tug on the miniscope while attached to the base plate while watching the imaging of the the brain. Moving the miniscope should not cause movement of the image on the video.<br />
*Make sure to have enough slack on the cable so the mice can move freely but not so much that the mice can chew on the cable. If you're not using a commutator, attend to the possible twisting of the cables while the animals are moving.<br />
*Check to see that the write speed of the computer exceeds the FPS collected. <br />
*After the trial, visually inspect that data folder to make sure the data was correctly collected.<br />
*Back up your data!!!</div>Chrisl1026http://miniscope.org/index.php?title=Animal_Behavior_Guide&diff=1551Animal Behavior Guide2016-11-16T01:12:04Z<p>Chrisl1026: /* Acclimation to experimenters/Handling */</p>
<hr />
<div>Here are some tips to for using the miniscopes to run experiments in freely moving mice.<br />
<br />
<br />
== Experimental timeline ==<br />
Plan you experiments schedule accordingly. Here is a typical timeline for getting mice ready for experiments in freely moving mice.<br />
<br />
:Week 1: virus injection surgery<br />
:Week 2: GRIN lens implant surgery<br />
:Week 3: recovery<br />
:Week 4: base plate surgery + acclimation to experimenters/handling<br />
:Week 5: habituation to wearing miniscopes & setting imaging focal plane<br />
:Week 6: begin experiment!<br />
<br />
== Acclimation to experimenters/Handling ==<br />
We typically wait at least a day after base plate surgery to begin handling. We recommend that mice become well acclimated to the experimenters prior to mounting miniscopes on awake moving mice. It is important that the mice feel comfortable with the experimenters as stress during the mounting of the miniscope during the experiment can cause adverse effects for the mice and thus, experiment. After gently handling the animals a few times, acclimate the mice to getting something placed on and off their heads by gently taking the protective caps on and off their base plate (that is cemented on the mice's heads). The experimenters should be able to quickly remove the cap without much struggle from the animal before continuing onto the habituation phase.<br />
<br />
<br clear=all><br />
<br />
{{#ev:youtube|https://youtu.be/RjH8wx-_ebM|640|center}}<br />
<br />
<br clear=all><br />
<br />
{{#ev:youtube|https://youtu.be/RjH8wx-_ebM|640|center}}<br />
<br />
<br clear=all><br />
<br />
== Habituation to wearing miniscopes ==<br />
Once the mice (and experimenters) are comfortable with the cap removal, habituate the mice to wearing the miniscopes while freely moving. To habituate, one experimenter should calmly hold the mouse and remove the cap, while the second experimenter replaces the cap with the miniscope. The miniscope should easily snap into place with the magnets. The second experimenter should gently hold the sides of the base plate with one hand while securing the miniscope with the set screw with the other hand. Especially during the first time, the animal may struggle. If the animal is struggling too much, let go and calm the animal. If the animal resists and struggles TOO much while you are holding onto the base plate, it can lead to the dental cement holding the base plate to come off the skull. When the animal is calm, try to hold the base plate and insert the set screw to secure the miniscope again. Once the miniscope is secured, let the animal explore your habituation environment of choice. We typically just let them move freely around in their home cage (even if there are other mice in the cage). Depending on your experimental demands, you will need to habituate to different levels of movement. If the experiment only requires mice to walk around and explore, we recommend habituating for at least 3 days for 10 minutes each day. If the experiment requires the mice to run swiftly (e.g. down a linear track), then we recommend habituating for at least 5 days for 10-30 minutes each day.<br />
<br />
== Setting imaging focal plane & LED intensity ==<br />
We recommend setting the focal plane of imaging and LED intensity prior to the experimental day. This can be done during the habituation sessions. Move the focus slider along the neck of the body of the miniscope until you've reached your optimal focal plane of the brain. To set the focal plane, tighten the set screw into the focus slider until the focus slider is secure. Be careful of over-tightening, as the screw can make indents into to the plastic material of the neck of the body. Once the focal plane is set, do not change it for the duration of your experiment if you need to image the same set of cells across sessions. Typically, we have found that for some mice, they can use the same miniscope set at a certain focal plane. For other mice, they need the miniscope to be set to a different height.<br />
<br />
During this session, you may also want to find the optimal level of LED intensity for imaging. Make sure that the there is enough blue light to see both the firing of the cells as well as some blood vessels to be used as landmarks to align frames. We recommend taking a video during the habituation session with your settings and analyzing the video to make sure the settings are optimal prior to the start of the experiment. Also, take a snapshot of the brain that you will use to reference for all future recording sessions. That way, you can begin your experiment with confidence and ease!<br />
<br />
== Starting an experiment ==<br />
Here is an example checklist that can be used.<br />
<br />
*Check if there is enough space on the hard drive for the data files that will be collected throughout the experiment. If you are using a behavioral camera, don't forget to include that as well.<br />
*Check and clean the imaging sensor before screwing it on the focus slider. If there is dust or dirt, use lens paper and alcohol to gently clean the sensor. This typically only needs to be done once prior to securing onto the focus slide. If the imaging sensor is being removed between sessions, make sure to screw it back onto the focus slide in the same orientation!<br />
*Connect the miniscope cable with the DAQ, connect the USB 3.0 cable with the DAQ and computer, and open the Miniscope Control application on the computer.<br />
*Make sure the 3 lights on the DAQ turn on and the red light on the miniscope turns on.<br />
*Make sure that the miniscope is flush against the base plate or as much as possible and the miniscope is not tilted and the set screw is not in the way.<br />
*Compare the live image you see with the snapshot of the same region you took during the habituation. Try to match it up as best as possible. The blood vessels, contrast, brightness should all look similar. If it doesn't then troubleshoot as to why it is different.<br />
*Gently tighten the set screw to secure the miniscope onto the base plate. Stop once the screw makes contact with the plastic of the miniscope. Over tightening will make an indentation into the plastic body and cause damage. To check if the miniscope is secure, GENTLY wiggle and tug on the miniscope while attached to the base plate while watching the imaging of the the brain. Moving the miniscope should not cause movement of the image on the video.<br />
*Make sure to have enough slack on the cable so the mice can move freely but not so much that the mice can chew on the cable. If you're not using a commutator, attend to the possible twisting of the cables while the animals are moving.<br />
*Check to see that the write speed of the computer exceeds the FPS collected. <br />
*After the trial, visually inspect that data folder to make sure the data was correctly collected.<br />
*Back up your data!!!</div>Chrisl1026http://miniscope.org/index.php?title=Surgery_Protocol&diff=1550Surgery Protocol2016-11-16T01:04:25Z<p>Chrisl1026: /* Baseplating */</p>
<hr />
<div>The following is a basic outline of our procedure for implanting a GRIN lens above the hippocampus for CA1 imaging. For videos of surgery and baseplating, see the [[Online Workshop]]. Also, see the recent Nature Protocols paper from the Stuber Lab ([http://www.nature.com/nprot/journal/v11/n3/full/nprot.2016.021.html link]) that has a very detailed description of the surgery. <br />
<br />
In order to image, animals will need to undergo three surgical procedures: a virus injection, a GRIN lens implantation, and baseplate attachment.<br />
<br />
== Virus Injection ==<br />
Before implanting the GRIN lens, you will need to inject a fluorescent indicator such as GCaMP6. We have generally used AAV1.Syn.GCaMP6f.WPRE.SV40 from Penn Vector (Item number AV-1-PV2822) and had great success in dorsal CA1. You may also be able to use GCaMP6 transgenic mice, but we have not tested whether any of the newer lines are bright enough to be imaged.<br />
<br />
== GRIN Lens Implantation==<br />
<br />
=== Basic Equipment Needed ===<br />
You will only need basic surgical equipment to perform GRIN lens implantation surgeries. This includes a mouse stereotax, an isoflurane vaporizor, surgical heat pad, stereo surgical microscope, light source, dental drill, and bead sterilizer. For recommended equipment, see the master [https://docs.google.com/spreadsheets/d/12H71DU2QX8d7efUE4yNuikBEiIzKaXjYqdc0A-oLNSw/edit?usp=sharing parts list].<br />
<br />
You will need the following basic tools and supplies: fine forceps, blunt forceps, fine scissors, scalpel and blade, small skull screws, drill bits, cyanoacrylate glue, dental cement, Kwik-Sil, and [[cortex buffer]]<br />
<br />
=== Aspirator and Lens Holder ===<br />
You will need an aspirator to remove cortex above the hippocampus. This can be built very simply with a vacuum line (or pump), a liquid trap, 1 ml syringe (with a hole to control suction), blunt needles, tubing and connectors, and hot glue.<br />
[[File:Aspirator.PNG|center|500px]]<br />
<br />
You can easily build a GRIN lens holder by using a suction system to hold the lens in place as it is dropped into the brain for implantation. To build, start with two 1ml micropipette tips. Cut the first so that the tip is just larger than the diameter of the GRIN lens you will be using. Cut the second tip so that the end is just smaller than the size of the GRIN lens. Connect this end to a 1ml syringe and connect to a vacuum line. This will then hold the lens in place during implantation. You can then use tape to attach this holder to a stereotax. Another solution we have used is a basic drill chuck, but this is only good for large (2mm) lenses.<br />
[[File:LensHolder.PNG|center|500px]]<br />
<br />
=== Skull Preparation ===<br />
You must first prepare the skull for implantation. We recommend shaving the head area and sterilizing with 3 alternating betadine and ethanol scrubs. Next, remove the scalp with scissors and clean the skull with hydrogen peroxide and saline. Scrape and score the skull to increase the bond with the glue. We also detach the neck muscle from the skull in order to reduce pull on the skull and prevent muscle growth that can make the implant less stable. Finally we insert a skull screw on the opposite side of the skull to enhance the stability of the implant.<br />
<br />
=== Craniotomy and Aspiration ===<br />
Using a stereotax, align the skull and find the location for the implant. Using a drill, make 4 guide holes to outline the lens placement. For CA1, we recommend offsetting the lens 0.5mm to the medial side of the virus injection to prevent imaging any damage induced by the virus injection. Connect the guide holes to create a circular craniotomy and peel off the circular skull fragment. Do not worry about blood as the top layers of cortex will be removed by aspiration. Make absolutely sure that the craniotomy is at least the size of your GRIN lens! Cover with cortex buffer and scrape around the outside of the craniotomy to remove any excess bone or dura. <br />
<br />
While keeping a constant flow of cortex buffer use the aspirator to slowly remove the cortex above the hippocampus. Continue removing cortex until you reach the white striations of the corpus callosum. Very slowly and carefully remove the horizontal white striations of the corpus callosum until you reach the striations that go vertical. When you reach these striations, stop aspirating and clean up the sides so that the lens can be placed into the brain. Continue washing with cortex buffer until all bleeding stops and keep the brain constantly flushed with cortex buffer.<br />
<br />
=== Lens implantation ===<br />
Attach GRIN lens to lens holder on stereotax. Align the lens with the top of the skull at the most posterior part of the craniotomy and quickly insert the lens 1.35 mm below the top of the skull. Remove any excess liquid with the aspirator. Connect the outside of the GRIN lens with the skull and skull screw using cyanoacrylate glue and let fully dry or use a glue accelerator. Remove the lens holder by simply removing the vacuum suction (you can just pinch the tubing) and withdrawing the holder. Next, cover the entire skull with cyanoacrylate glue, and then cover with dental cement. Let dry completely and cover with Kwik-Sil to protect the lens. Remove from stereotax and allow the animal to recover. Give the animal amoxicillin (or equivalent) for 7 days, and daily injections of carprofen and dexamethasone for 7 days.<br />
<br />
=== Important tips to remember ===<br />
*Try to keep the surgery as sterile as possible. Infections will destroy imaging and the ability to keep the same cells over many days.<br />
<br />
*Try to keep isoflurane as low as possible. Surgeries can take 2-6 hours depending on experience.<br />
<br />
*Aspirations will take a lot of practice. Go slow at first.<br />
<br />
*Depth will be trial and error. If the first few animals don't give good images, check the depth using histology and adjust accordingly.<br />
<br />
<br />
Please feel free to add any tips that you have!<br />
<br />
== Baseplating Protocol ==<br />
<br />
<br />
=== Basic Equipment needed ===<br />
[[File:bp1.png|right|200px|thumb|Put a stack of magnets on a reference Miniscope and mark the top magnet to keep track of its polarity]]<br />
[[File:bp2.png|right|200px|thumb|Magnetically attach the top magnet to screwdriver, coat with cyanoacrylate glue on the edge, and push through hole on baseplate]]<br />
[[File:bp3.png|right|200px|thumb|Repeat for remaining two magnets; ensure top of baseplate is completely flat]]<br />
[[File:bp4.png|right|200px|thumb|Coat bottom of baseplate with cyanoacrylate glue]]<br />
[[File:bp5.png|right|200px|thumb|Score bottom and edges of baseplate with a dental drill]]<br />
You will only need basic surgical equipment to baseplate. This includes a mouse stereotax, an isoflurane vaporizer, surgical heat pad, stereo surgical microscope, light source, and a dental drill. <br />
<br />
You will also need cyanoacrylate glue, fine forceps, double distilled water, lens paper, and dental cement<br />
<br />
=== Baseplate Preparation ===<br />
Before affixing the baseplate on an animal, you will need to prepare the baseplate. <br />
The first step is to insert magnets into the appropriate holes on the baseplate. To ensure that the polarity of the magnets is correct throughout the baseplate preparation, use a reference Miniscope to hold onto the magnets. Mark the top magnet with a sharpie, as seen in the picture to the right. Next, put the top magnet on the head of a screwdriver with the marked side facing away from the screwdriver. Apply cyanoacrylate glue on the edge of the magnet, then push the magnet through the appropriate hole. Ensure that the magnet goes through the hole completely and the top side of the baseplate is flat, as the quality of imaging relies on the Miniscope sitting flush on the baseplate. Repeat this procedure for the next 2 magnets. Finally, put a coat of cyanoacrylate glue on the bottom side of the baseplate and leave the baseplate to dry.<br />
Once the baseplate has dried, score the bottom and the sides of the baseplate using a dental drill to allow the dental glue to adhere the baseplate and skull well.<br />
<br />
<br />
{{#ev:youtube|https://youtu.be/z_32O2XoYI4|640|center}}<br />
<br />
=== Cap Preparation ===<br />
Similar to the baseplate preparation, use a reference Miniscope to hold onto the magnets. Mark the top magnet with a sharpie, and put the magnet on the head of a screwdriver with the marked side facing toward the screwdriver. Coat the magnet with cyanoacrylate glue and push the magnet into the appropriate hole. Repeat for the remaining two magnets.<br />
=== Baseplating ===<br />
Connect a Miniscope to a DAQ and initialize the Miniscope software on a computer. Attach your prepared baseplate onto the bottom of the Miniscope using a set screw.<br />
<br />
Using a stereotax and isoflurane vaporizer, anesthetize the animal and fix ear bars until the animal is stable. Carefully remove the Kwik-Sil off the top of the animal’s GRIN lens with blunt forceps being sure not to scratch the lens. Under a stereo surgical microscope, check to see if there is debris on the top of the GRIN lens. If there is, dampen a sheet of lens paper using double distilled water. Wipe the top of the GRIN lens using fine forceps and lens paper, being sure to only wipe laterally and not to apply direct downward pressure.<br />
<br />
After the top of the GRIN lens is clean, explore the field of view with the Miniscope. Adjust the LED brightness and focus accordingly. We advise that you use maximum gain and low LED excitation as to not photobleach the brain during baseplating. After finding your ideal view, practice taking the Miniscope off and on and finding the same view. <br />
<br />
When you feel comfortable repeatedly finding the view with the Miniscope, apply dental cement in a “C” pattern around the GRIN lens, making sure not to touch the lens itself. It will be easier if your first application of dental cement is more viscous. Put the Miniscope on top of the lens and find your ideal view. Hold the Miniscope steadily as the first application of dental cement dries, making sure that your view does not change. <br />
After the first application of dental cement dries, mix more dental cement and apply it on the posterior edge of the baseplate, making sure it seals to the skull. At this point, you can also build up dental cement along the wall of the baseplate on the animal’s right side. Be sure to avoid applying dental cement onto the camera itself and adhering it to the baseplate. Once the dental cement dries and you feel confident that the baseplate will no longer shift, carefully remove the Miniscope from the baseplate by unscrewing the set screw and lifting it off. <br />
<br />
Apply dental cement to the anterior edge of the baseplate and fill in the gaps between the baseplate and skull on the animal’s left side. Allow the dental cement to dry and attach the plastic cap onto the top of the baseplate with a set screw.<br />
<br />
{{#ev:youtube|https://youtu.be/GoDJGfqO3po|640|center}}</div>Chrisl1026http://miniscope.org/index.php?title=Surgery_Protocol&diff=1549Surgery Protocol2016-11-16T01:03:30Z<p>Chrisl1026: /* Baseplating */</p>
<hr />
<div>The following is a basic outline of our procedure for implanting a GRIN lens above the hippocampus for CA1 imaging. For videos of surgery and baseplating, see the [[Online Workshop]]. Also, see the recent Nature Protocols paper from the Stuber Lab ([http://www.nature.com/nprot/journal/v11/n3/full/nprot.2016.021.html link]) that has a very detailed description of the surgery. <br />
<br />
In order to image, animals will need to undergo three surgical procedures: a virus injection, a GRIN lens implantation, and baseplate attachment.<br />
<br />
== Virus Injection ==<br />
Before implanting the GRIN lens, you will need to inject a fluorescent indicator such as GCaMP6. We have generally used AAV1.Syn.GCaMP6f.WPRE.SV40 from Penn Vector (Item number AV-1-PV2822) and had great success in dorsal CA1. You may also be able to use GCaMP6 transgenic mice, but we have not tested whether any of the newer lines are bright enough to be imaged.<br />
<br />
== GRIN Lens Implantation==<br />
<br />
=== Basic Equipment Needed ===<br />
You will only need basic surgical equipment to perform GRIN lens implantation surgeries. This includes a mouse stereotax, an isoflurane vaporizor, surgical heat pad, stereo surgical microscope, light source, dental drill, and bead sterilizer. For recommended equipment, see the master [https://docs.google.com/spreadsheets/d/12H71DU2QX8d7efUE4yNuikBEiIzKaXjYqdc0A-oLNSw/edit?usp=sharing parts list].<br />
<br />
You will need the following basic tools and supplies: fine forceps, blunt forceps, fine scissors, scalpel and blade, small skull screws, drill bits, cyanoacrylate glue, dental cement, Kwik-Sil, and [[cortex buffer]]<br />
<br />
=== Aspirator and Lens Holder ===<br />
You will need an aspirator to remove cortex above the hippocampus. This can be built very simply with a vacuum line (or pump), a liquid trap, 1 ml syringe (with a hole to control suction), blunt needles, tubing and connectors, and hot glue.<br />
[[File:Aspirator.PNG|center|500px]]<br />
<br />
You can easily build a GRIN lens holder by using a suction system to hold the lens in place as it is dropped into the brain for implantation. To build, start with two 1ml micropipette tips. Cut the first so that the tip is just larger than the diameter of the GRIN lens you will be using. Cut the second tip so that the end is just smaller than the size of the GRIN lens. Connect this end to a 1ml syringe and connect to a vacuum line. This will then hold the lens in place during implantation. You can then use tape to attach this holder to a stereotax. Another solution we have used is a basic drill chuck, but this is only good for large (2mm) lenses.<br />
[[File:LensHolder.PNG|center|500px]]<br />
<br />
=== Skull Preparation ===<br />
You must first prepare the skull for implantation. We recommend shaving the head area and sterilizing with 3 alternating betadine and ethanol scrubs. Next, remove the scalp with scissors and clean the skull with hydrogen peroxide and saline. Scrape and score the skull to increase the bond with the glue. We also detach the neck muscle from the skull in order to reduce pull on the skull and prevent muscle growth that can make the implant less stable. Finally we insert a skull screw on the opposite side of the skull to enhance the stability of the implant.<br />
<br />
=== Craniotomy and Aspiration ===<br />
Using a stereotax, align the skull and find the location for the implant. Using a drill, make 4 guide holes to outline the lens placement. For CA1, we recommend offsetting the lens 0.5mm to the medial side of the virus injection to prevent imaging any damage induced by the virus injection. Connect the guide holes to create a circular craniotomy and peel off the circular skull fragment. Do not worry about blood as the top layers of cortex will be removed by aspiration. Make absolutely sure that the craniotomy is at least the size of your GRIN lens! Cover with cortex buffer and scrape around the outside of the craniotomy to remove any excess bone or dura. <br />
<br />
While keeping a constant flow of cortex buffer use the aspirator to slowly remove the cortex above the hippocampus. Continue removing cortex until you reach the white striations of the corpus callosum. Very slowly and carefully remove the horizontal white striations of the corpus callosum until you reach the striations that go vertical. When you reach these striations, stop aspirating and clean up the sides so that the lens can be placed into the brain. Continue washing with cortex buffer until all bleeding stops and keep the brain constantly flushed with cortex buffer.<br />
<br />
=== Lens implantation ===<br />
Attach GRIN lens to lens holder on stereotax. Align the lens with the top of the skull at the most posterior part of the craniotomy and quickly insert the lens 1.35 mm below the top of the skull. Remove any excess liquid with the aspirator. Connect the outside of the GRIN lens with the skull and skull screw using cyanoacrylate glue and let fully dry or use a glue accelerator. Remove the lens holder by simply removing the vacuum suction (you can just pinch the tubing) and withdrawing the holder. Next, cover the entire skull with cyanoacrylate glue, and then cover with dental cement. Let dry completely and cover with Kwik-Sil to protect the lens. Remove from stereotax and allow the animal to recover. Give the animal amoxicillin (or equivalent) for 7 days, and daily injections of carprofen and dexamethasone for 7 days.<br />
<br />
=== Important tips to remember ===<br />
*Try to keep the surgery as sterile as possible. Infections will destroy imaging and the ability to keep the same cells over many days.<br />
<br />
*Try to keep isoflurane as low as possible. Surgeries can take 2-6 hours depending on experience.<br />
<br />
*Aspirations will take a lot of practice. Go slow at first.<br />
<br />
*Depth will be trial and error. If the first few animals don't give good images, check the depth using histology and adjust accordingly.<br />
<br />
<br />
Please feel free to add any tips that you have!<br />
<br />
== Baseplating Protocol ==<br />
<br />
<br />
=== Basic Equipment needed ===<br />
[[File:bp1.png|right|200px|thumb|Put a stack of magnets on a reference Miniscope and mark the top magnet to keep track of its polarity]]<br />
[[File:bp2.png|right|200px|thumb|Magnetically attach the top magnet to screwdriver, coat with cyanoacrylate glue on the edge, and push through hole on baseplate]]<br />
[[File:bp3.png|right|200px|thumb|Repeat for remaining two magnets; ensure top of baseplate is completely flat]]<br />
[[File:bp4.png|right|200px|thumb|Coat bottom of baseplate with cyanoacrylate glue]]<br />
[[File:bp5.png|right|200px|thumb|Score bottom and edges of baseplate with a dental drill]]<br />
You will only need basic surgical equipment to baseplate. This includes a mouse stereotax, an isoflurane vaporizer, surgical heat pad, stereo surgical microscope, light source, and a dental drill. <br />
<br />
You will also need cyanoacrylate glue, fine forceps, double distilled water, lens paper, and dental cement<br />
<br />
=== Baseplate Preparation ===<br />
Before affixing the baseplate on an animal, you will need to prepare the baseplate. <br />
The first step is to insert magnets into the appropriate holes on the baseplate. To ensure that the polarity of the magnets is correct throughout the baseplate preparation, use a reference Miniscope to hold onto the magnets. Mark the top magnet with a sharpie, as seen in the picture to the right. Next, put the top magnet on the head of a screwdriver with the marked side facing away from the screwdriver. Apply cyanoacrylate glue on the edge of the magnet, then push the magnet through the appropriate hole. Ensure that the magnet goes through the hole completely and the top side of the baseplate is flat, as the quality of imaging relies on the Miniscope sitting flush on the baseplate. Repeat this procedure for the next 2 magnets. Finally, put a coat of cyanoacrylate glue on the bottom side of the baseplate and leave the baseplate to dry.<br />
Once the baseplate has dried, score the bottom and the sides of the baseplate using a dental drill to allow the dental glue to adhere the baseplate and skull well.<br />
<br />
<br />
{{#ev:youtube|https://youtu.be/z_32O2XoYI4|640|center}}<br />
<br />
=== Cap Preparation ===<br />
Similar to the baseplate preparation, use a reference Miniscope to hold onto the magnets. Mark the top magnet with a sharpie, and put the magnet on the head of a screwdriver with the marked side facing toward the screwdriver. Coat the magnet with cyanoacrylate glue and push the magnet into the appropriate hole. Repeat for the remaining two magnets.<br />
=== Baseplating ===<br />
Connect a Miniscope to a DAQ and initialize the Miniscope software on a computer. Attach your prepared baseplate onto the bottom of the Miniscope using a set screw.<br />
<br />
Using a stereotax and isoflurane vaporizer, anesthetize the animal and fix ear bars until the animal is stable. Carefully remove the Kwik-Sil off the top of the animal’s GRIN lens with blunt forceps being sure not to scratch the lens. Under a stereo surgical microscope, check to see if there is debris on the top of the GRIN lens. If there is, dampen a sheet of lens paper using double distilled water. Wipe the top of the GRIN lens using fine forceps and lens paper, being sure to only wipe laterally and not to apply direct downward pressure.<br />
<br />
After the top of the GRIN lens is clean, explore the field of view with the Miniscope. Adjust the LED brightness and focus accordingly. We advise that you use maximum gain and low LED excitation as to not photobleach the brain during baseplating. After finding your ideal view, practice taking the Miniscope off and on and finding the same view. <br />
<br />
When you feel comfortable repeatedly finding the view with the Miniscope, apply dental cement in a “C” pattern around the GRIN lens, making sure not to touch the lens itself. It will be easier if your first application of dental cement is more viscous. Put the Miniscope on top of the lens and find your ideal view. Hold the Miniscope steadily as the first application of dental cement dries, making sure that your view does not change. <br />
After the first application of dental cement dries, mix more dental cement and apply it on the posterior edge of the baseplate, making sure it seals to the skull. At this point, you can also build up dental cement along the wall of the baseplate on the animal’s right side. Be sure to avoid applying dental cement onto the camera itself and adhering it to the baseplate. Once the dental cement dries and you feel confident that the baseplate will no longer shift, carefully remove the Miniscope from the baseplate by unscrewing the set screw and lifting it off. <br />
<br />
Apply dental cement to the anterior edge of the baseplate and fill in the gaps between the baseplate and skull on the animal’s left side. Allow the dental cement to dry and attach the plastic cap onto the top of the baseplate with a set screw.<br />
<br />
https://youtu.be/GoDJGfqO3po</div>Chrisl1026http://miniscope.org/index.php?title=Surgery_Protocol&diff=1548Surgery Protocol2016-11-16T01:02:51Z<p>Chrisl1026: /* Baseplate Preparation */</p>
<hr />
<div>The following is a basic outline of our procedure for implanting a GRIN lens above the hippocampus for CA1 imaging. For videos of surgery and baseplating, see the [[Online Workshop]]. Also, see the recent Nature Protocols paper from the Stuber Lab ([http://www.nature.com/nprot/journal/v11/n3/full/nprot.2016.021.html link]) that has a very detailed description of the surgery. <br />
<br />
In order to image, animals will need to undergo three surgical procedures: a virus injection, a GRIN lens implantation, and baseplate attachment.<br />
<br />
== Virus Injection ==<br />
Before implanting the GRIN lens, you will need to inject a fluorescent indicator such as GCaMP6. We have generally used AAV1.Syn.GCaMP6f.WPRE.SV40 from Penn Vector (Item number AV-1-PV2822) and had great success in dorsal CA1. You may also be able to use GCaMP6 transgenic mice, but we have not tested whether any of the newer lines are bright enough to be imaged.<br />
<br />
== GRIN Lens Implantation==<br />
<br />
=== Basic Equipment Needed ===<br />
You will only need basic surgical equipment to perform GRIN lens implantation surgeries. This includes a mouse stereotax, an isoflurane vaporizor, surgical heat pad, stereo surgical microscope, light source, dental drill, and bead sterilizer. For recommended equipment, see the master [https://docs.google.com/spreadsheets/d/12H71DU2QX8d7efUE4yNuikBEiIzKaXjYqdc0A-oLNSw/edit?usp=sharing parts list].<br />
<br />
You will need the following basic tools and supplies: fine forceps, blunt forceps, fine scissors, scalpel and blade, small skull screws, drill bits, cyanoacrylate glue, dental cement, Kwik-Sil, and [[cortex buffer]]<br />
<br />
=== Aspirator and Lens Holder ===<br />
You will need an aspirator to remove cortex above the hippocampus. This can be built very simply with a vacuum line (or pump), a liquid trap, 1 ml syringe (with a hole to control suction), blunt needles, tubing and connectors, and hot glue.<br />
[[File:Aspirator.PNG|center|500px]]<br />
<br />
You can easily build a GRIN lens holder by using a suction system to hold the lens in place as it is dropped into the brain for implantation. To build, start with two 1ml micropipette tips. Cut the first so that the tip is just larger than the diameter of the GRIN lens you will be using. Cut the second tip so that the end is just smaller than the size of the GRIN lens. Connect this end to a 1ml syringe and connect to a vacuum line. This will then hold the lens in place during implantation. You can then use tape to attach this holder to a stereotax. Another solution we have used is a basic drill chuck, but this is only good for large (2mm) lenses.<br />
[[File:LensHolder.PNG|center|500px]]<br />
<br />
=== Skull Preparation ===<br />
You must first prepare the skull for implantation. We recommend shaving the head area and sterilizing with 3 alternating betadine and ethanol scrubs. Next, remove the scalp with scissors and clean the skull with hydrogen peroxide and saline. Scrape and score the skull to increase the bond with the glue. We also detach the neck muscle from the skull in order to reduce pull on the skull and prevent muscle growth that can make the implant less stable. Finally we insert a skull screw on the opposite side of the skull to enhance the stability of the implant.<br />
<br />
=== Craniotomy and Aspiration ===<br />
Using a stereotax, align the skull and find the location for the implant. Using a drill, make 4 guide holes to outline the lens placement. For CA1, we recommend offsetting the lens 0.5mm to the medial side of the virus injection to prevent imaging any damage induced by the virus injection. Connect the guide holes to create a circular craniotomy and peel off the circular skull fragment. Do not worry about blood as the top layers of cortex will be removed by aspiration. Make absolutely sure that the craniotomy is at least the size of your GRIN lens! Cover with cortex buffer and scrape around the outside of the craniotomy to remove any excess bone or dura. <br />
<br />
While keeping a constant flow of cortex buffer use the aspirator to slowly remove the cortex above the hippocampus. Continue removing cortex until you reach the white striations of the corpus callosum. Very slowly and carefully remove the horizontal white striations of the corpus callosum until you reach the striations that go vertical. When you reach these striations, stop aspirating and clean up the sides so that the lens can be placed into the brain. Continue washing with cortex buffer until all bleeding stops and keep the brain constantly flushed with cortex buffer.<br />
<br />
=== Lens implantation ===<br />
Attach GRIN lens to lens holder on stereotax. Align the lens with the top of the skull at the most posterior part of the craniotomy and quickly insert the lens 1.35 mm below the top of the skull. Remove any excess liquid with the aspirator. Connect the outside of the GRIN lens with the skull and skull screw using cyanoacrylate glue and let fully dry or use a glue accelerator. Remove the lens holder by simply removing the vacuum suction (you can just pinch the tubing) and withdrawing the holder. Next, cover the entire skull with cyanoacrylate glue, and then cover with dental cement. Let dry completely and cover with Kwik-Sil to protect the lens. Remove from stereotax and allow the animal to recover. Give the animal amoxicillin (or equivalent) for 7 days, and daily injections of carprofen and dexamethasone for 7 days.<br />
<br />
=== Important tips to remember ===<br />
*Try to keep the surgery as sterile as possible. Infections will destroy imaging and the ability to keep the same cells over many days.<br />
<br />
*Try to keep isoflurane as low as possible. Surgeries can take 2-6 hours depending on experience.<br />
<br />
*Aspirations will take a lot of practice. Go slow at first.<br />
<br />
*Depth will be trial and error. If the first few animals don't give good images, check the depth using histology and adjust accordingly.<br />
<br />
<br />
Please feel free to add any tips that you have!<br />
<br />
== Baseplating Protocol ==<br />
<br />
<br />
=== Basic Equipment needed ===<br />
[[File:bp1.png|right|200px|thumb|Put a stack of magnets on a reference Miniscope and mark the top magnet to keep track of its polarity]]<br />
[[File:bp2.png|right|200px|thumb|Magnetically attach the top magnet to screwdriver, coat with cyanoacrylate glue on the edge, and push through hole on baseplate]]<br />
[[File:bp3.png|right|200px|thumb|Repeat for remaining two magnets; ensure top of baseplate is completely flat]]<br />
[[File:bp4.png|right|200px|thumb|Coat bottom of baseplate with cyanoacrylate glue]]<br />
[[File:bp5.png|right|200px|thumb|Score bottom and edges of baseplate with a dental drill]]<br />
You will only need basic surgical equipment to baseplate. This includes a mouse stereotax, an isoflurane vaporizer, surgical heat pad, stereo surgical microscope, light source, and a dental drill. <br />
<br />
You will also need cyanoacrylate glue, fine forceps, double distilled water, lens paper, and dental cement<br />
<br />
=== Baseplate Preparation ===<br />
Before affixing the baseplate on an animal, you will need to prepare the baseplate. <br />
The first step is to insert magnets into the appropriate holes on the baseplate. To ensure that the polarity of the magnets is correct throughout the baseplate preparation, use a reference Miniscope to hold onto the magnets. Mark the top magnet with a sharpie, as seen in the picture to the right. Next, put the top magnet on the head of a screwdriver with the marked side facing away from the screwdriver. Apply cyanoacrylate glue on the edge of the magnet, then push the magnet through the appropriate hole. Ensure that the magnet goes through the hole completely and the top side of the baseplate is flat, as the quality of imaging relies on the Miniscope sitting flush on the baseplate. Repeat this procedure for the next 2 magnets. Finally, put a coat of cyanoacrylate glue on the bottom side of the baseplate and leave the baseplate to dry.<br />
Once the baseplate has dried, score the bottom and the sides of the baseplate using a dental drill to allow the dental glue to adhere the baseplate and skull well.<br />
<br />
<br />
{{#ev:youtube|https://youtu.be/z_32O2XoYI4|640|center}}<br />
<br />
=== Cap Preparation ===<br />
Similar to the baseplate preparation, use a reference Miniscope to hold onto the magnets. Mark the top magnet with a sharpie, and put the magnet on the head of a screwdriver with the marked side facing toward the screwdriver. Coat the magnet with cyanoacrylate glue and push the magnet into the appropriate hole. Repeat for the remaining two magnets.<br />
=== Baseplating ===<br />
Connect a Miniscope to a DAQ and initialize the Miniscope software on a computer. Attach your prepared baseplate onto the bottom of the Miniscope using a set screw.<br />
<br />
Using a stereotax and isoflurane vaporizer, anesthetize the animal and fix ear bars until the animal is stable. Carefully remove the Kwik-Sil off the top of the animal’s GRIN lens with blunt forceps being sure not to scratch the lens. Under a stereo surgical microscope, check to see if there is debris on the top of the GRIN lens. If there is, dampen a sheet of lens paper using double distilled water. Wipe the top of the GRIN lens using fine forceps and lens paper, being sure to only wipe laterally and not to apply direct downward pressure.<br />
<br />
After the top of the GRIN lens is clean, explore the field of view with the Miniscope. Adjust the LED brightness and focus accordingly. We advise that you use maximum gain and low LED excitation as to not photobleach the brain during baseplating. After finding your ideal view, practice taking the Miniscope off and on and finding the same view. <br />
<br />
When you feel comfortable repeatedly finding the view with the Miniscope, apply dental cement in a “C” pattern around the GRIN lens, making sure not to touch the lens itself. It will be easier if your first application of dental cement is more viscous. Put the Miniscope on top of the lens and find your ideal view. Hold the Miniscope steadily as the first application of dental cement dries, making sure that your view does not change. <br />
After the first application of dental cement dries, mix more dental cement and apply it on the posterior edge of the baseplate, making sure it seals to the skull. At this point, you can also build up dental cement along the wall of the baseplate on the animal’s right side. Be sure to avoid applying dental cement onto the camera itself and adhering it to the baseplate. Once the dental cement dries and you feel confident that the baseplate will no longer shift, carefully remove the Miniscope from the baseplate by unscrewing the set screw and lifting it off. <br />
<br />
Apply dental cement to the anterior edge of the baseplate and fill in the gaps between the baseplate and skull on the animal’s left side. Allow the dental cement to dry and attach the plastic cap onto the top of the baseplate with a set screw.</div>Chrisl1026http://miniscope.org/index.php?title=Surgery_Protocol&diff=1547Surgery Protocol2016-11-16T01:02:31Z<p>Chrisl1026: /* Baseplate Preparation */</p>
<hr />
<div>The following is a basic outline of our procedure for implanting a GRIN lens above the hippocampus for CA1 imaging. For videos of surgery and baseplating, see the [[Online Workshop]]. Also, see the recent Nature Protocols paper from the Stuber Lab ([http://www.nature.com/nprot/journal/v11/n3/full/nprot.2016.021.html link]) that has a very detailed description of the surgery. <br />
<br />
In order to image, animals will need to undergo three surgical procedures: a virus injection, a GRIN lens implantation, and baseplate attachment.<br />
<br />
== Virus Injection ==<br />
Before implanting the GRIN lens, you will need to inject a fluorescent indicator such as GCaMP6. We have generally used AAV1.Syn.GCaMP6f.WPRE.SV40 from Penn Vector (Item number AV-1-PV2822) and had great success in dorsal CA1. You may also be able to use GCaMP6 transgenic mice, but we have not tested whether any of the newer lines are bright enough to be imaged.<br />
<br />
== GRIN Lens Implantation==<br />
<br />
=== Basic Equipment Needed ===<br />
You will only need basic surgical equipment to perform GRIN lens implantation surgeries. This includes a mouse stereotax, an isoflurane vaporizor, surgical heat pad, stereo surgical microscope, light source, dental drill, and bead sterilizer. For recommended equipment, see the master [https://docs.google.com/spreadsheets/d/12H71DU2QX8d7efUE4yNuikBEiIzKaXjYqdc0A-oLNSw/edit?usp=sharing parts list].<br />
<br />
You will need the following basic tools and supplies: fine forceps, blunt forceps, fine scissors, scalpel and blade, small skull screws, drill bits, cyanoacrylate glue, dental cement, Kwik-Sil, and [[cortex buffer]]<br />
<br />
=== Aspirator and Lens Holder ===<br />
You will need an aspirator to remove cortex above the hippocampus. This can be built very simply with a vacuum line (or pump), a liquid trap, 1 ml syringe (with a hole to control suction), blunt needles, tubing and connectors, and hot glue.<br />
[[File:Aspirator.PNG|center|500px]]<br />
<br />
You can easily build a GRIN lens holder by using a suction system to hold the lens in place as it is dropped into the brain for implantation. To build, start with two 1ml micropipette tips. Cut the first so that the tip is just larger than the diameter of the GRIN lens you will be using. Cut the second tip so that the end is just smaller than the size of the GRIN lens. Connect this end to a 1ml syringe and connect to a vacuum line. This will then hold the lens in place during implantation. You can then use tape to attach this holder to a stereotax. Another solution we have used is a basic drill chuck, but this is only good for large (2mm) lenses.<br />
[[File:LensHolder.PNG|center|500px]]<br />
<br />
=== Skull Preparation ===<br />
You must first prepare the skull for implantation. We recommend shaving the head area and sterilizing with 3 alternating betadine and ethanol scrubs. Next, remove the scalp with scissors and clean the skull with hydrogen peroxide and saline. Scrape and score the skull to increase the bond with the glue. We also detach the neck muscle from the skull in order to reduce pull on the skull and prevent muscle growth that can make the implant less stable. Finally we insert a skull screw on the opposite side of the skull to enhance the stability of the implant.<br />
<br />
=== Craniotomy and Aspiration ===<br />
Using a stereotax, align the skull and find the location for the implant. Using a drill, make 4 guide holes to outline the lens placement. For CA1, we recommend offsetting the lens 0.5mm to the medial side of the virus injection to prevent imaging any damage induced by the virus injection. Connect the guide holes to create a circular craniotomy and peel off the circular skull fragment. Do not worry about blood as the top layers of cortex will be removed by aspiration. Make absolutely sure that the craniotomy is at least the size of your GRIN lens! Cover with cortex buffer and scrape around the outside of the craniotomy to remove any excess bone or dura. <br />
<br />
While keeping a constant flow of cortex buffer use the aspirator to slowly remove the cortex above the hippocampus. Continue removing cortex until you reach the white striations of the corpus callosum. Very slowly and carefully remove the horizontal white striations of the corpus callosum until you reach the striations that go vertical. When you reach these striations, stop aspirating and clean up the sides so that the lens can be placed into the brain. Continue washing with cortex buffer until all bleeding stops and keep the brain constantly flushed with cortex buffer.<br />
<br />
=== Lens implantation ===<br />
Attach GRIN lens to lens holder on stereotax. Align the lens with the top of the skull at the most posterior part of the craniotomy and quickly insert the lens 1.35 mm below the top of the skull. Remove any excess liquid with the aspirator. Connect the outside of the GRIN lens with the skull and skull screw using cyanoacrylate glue and let fully dry or use a glue accelerator. Remove the lens holder by simply removing the vacuum suction (you can just pinch the tubing) and withdrawing the holder. Next, cover the entire skull with cyanoacrylate glue, and then cover with dental cement. Let dry completely and cover with Kwik-Sil to protect the lens. Remove from stereotax and allow the animal to recover. Give the animal amoxicillin (or equivalent) for 7 days, and daily injections of carprofen and dexamethasone for 7 days.<br />
<br />
=== Important tips to remember ===<br />
*Try to keep the surgery as sterile as possible. Infections will destroy imaging and the ability to keep the same cells over many days.<br />
<br />
*Try to keep isoflurane as low as possible. Surgeries can take 2-6 hours depending on experience.<br />
<br />
*Aspirations will take a lot of practice. Go slow at first.<br />
<br />
*Depth will be trial and error. If the first few animals don't give good images, check the depth using histology and adjust accordingly.<br />
<br />
<br />
Please feel free to add any tips that you have!<br />
<br />
== Baseplating Protocol ==<br />
<br />
<br />
=== Basic Equipment needed ===<br />
[[File:bp1.png|right|200px|thumb|Put a stack of magnets on a reference Miniscope and mark the top magnet to keep track of its polarity]]<br />
[[File:bp2.png|right|200px|thumb|Magnetically attach the top magnet to screwdriver, coat with cyanoacrylate glue on the edge, and push through hole on baseplate]]<br />
[[File:bp3.png|right|200px|thumb|Repeat for remaining two magnets; ensure top of baseplate is completely flat]]<br />
[[File:bp4.png|right|200px|thumb|Coat bottom of baseplate with cyanoacrylate glue]]<br />
[[File:bp5.png|right|200px|thumb|Score bottom and edges of baseplate with a dental drill]]<br />
You will only need basic surgical equipment to baseplate. This includes a mouse stereotax, an isoflurane vaporizer, surgical heat pad, stereo surgical microscope, light source, and a dental drill. <br />
<br />
You will also need cyanoacrylate glue, fine forceps, double distilled water, lens paper, and dental cement<br />
<br />
=== Baseplate Preparation ===<br />
Before affixing the baseplate on an animal, you will need to prepare the baseplate. <br />
The first step is to insert magnets into the appropriate holes on the baseplate. To ensure that the polarity of the magnets is correct throughout the baseplate preparation, use a reference Miniscope to hold onto the magnets. Mark the top magnet with a sharpie, as seen in the picture to the right. Next, put the top magnet on the head of a screwdriver with the marked side facing away from the screwdriver. Apply cyanoacrylate glue on the edge of the magnet, then push the magnet through the appropriate hole. Ensure that the magnet goes through the hole completely and the top side of the baseplate is flat, as the quality of imaging relies on the Miniscope sitting flush on the baseplate. Repeat this procedure for the next 2 magnets. Finally, put a coat of cyanoacrylate glue on the bottom side of the baseplate and leave the baseplate to dry.<br />
Once the baseplate has dried, score the bottom and the sides of the baseplate using a dental drill to allow the dental glue to adhere the baseplate and skull well.<br />
<br />
<br clear=all><br />
<br />
{{#ev:youtube|https://youtu.be/z_32O2XoYI4|640|center}}<br />
<br />
<br clear=all><br />
<br />
=== Cap Preparation ===<br />
Similar to the baseplate preparation, use a reference Miniscope to hold onto the magnets. Mark the top magnet with a sharpie, and put the magnet on the head of a screwdriver with the marked side facing toward the screwdriver. Coat the magnet with cyanoacrylate glue and push the magnet into the appropriate hole. Repeat for the remaining two magnets.<br />
=== Baseplating ===<br />
Connect a Miniscope to a DAQ and initialize the Miniscope software on a computer. Attach your prepared baseplate onto the bottom of the Miniscope using a set screw.<br />
<br />
Using a stereotax and isoflurane vaporizer, anesthetize the animal and fix ear bars until the animal is stable. Carefully remove the Kwik-Sil off the top of the animal’s GRIN lens with blunt forceps being sure not to scratch the lens. Under a stereo surgical microscope, check to see if there is debris on the top of the GRIN lens. If there is, dampen a sheet of lens paper using double distilled water. Wipe the top of the GRIN lens using fine forceps and lens paper, being sure to only wipe laterally and not to apply direct downward pressure.<br />
<br />
After the top of the GRIN lens is clean, explore the field of view with the Miniscope. Adjust the LED brightness and focus accordingly. We advise that you use maximum gain and low LED excitation as to not photobleach the brain during baseplating. After finding your ideal view, practice taking the Miniscope off and on and finding the same view. <br />
<br />
When you feel comfortable repeatedly finding the view with the Miniscope, apply dental cement in a “C” pattern around the GRIN lens, making sure not to touch the lens itself. It will be easier if your first application of dental cement is more viscous. Put the Miniscope on top of the lens and find your ideal view. Hold the Miniscope steadily as the first application of dental cement dries, making sure that your view does not change. <br />
After the first application of dental cement dries, mix more dental cement and apply it on the posterior edge of the baseplate, making sure it seals to the skull. At this point, you can also build up dental cement along the wall of the baseplate on the animal’s right side. Be sure to avoid applying dental cement onto the camera itself and adhering it to the baseplate. Once the dental cement dries and you feel confident that the baseplate will no longer shift, carefully remove the Miniscope from the baseplate by unscrewing the set screw and lifting it off. <br />
<br />
Apply dental cement to the anterior edge of the baseplate and fill in the gaps between the baseplate and skull on the animal’s left side. Allow the dental cement to dry and attach the plastic cap onto the top of the baseplate with a set screw.</div>Chrisl1026http://miniscope.org/index.php?title=Imaging_With_Thin_GRIN_Lenses&diff=1546Imaging With Thin GRIN Lenses2016-11-14T19:29:30Z<p>Chrisl1026: /* Modification Overview */</p>
<hr />
<div>Want to image with thin GRIN lenses? Follow the simple steps below for modifying your scope to be able to image through 1mm diameter and thinner GRIN relay lenses. Also make sure you have read our [[GRIN Lens Information]] page.<br />
<br />
= Modification Overview =<br />
The Miniscope relies on a 0.25pitch, 2mm or 1.8mm diameter, objective GRIN lens for imaging. This lens can be implanted directly into the brain ([[Surgery Protocol]], [https://drive.google.com/file/d/0ByUbjrn9MxK0TWdxUVVjakF3cDQ/view?usp=sharing| Surgery and Baseplating Presentation]) but it can also be mounted into the scope to allow for imaging through thinner diameter GRIN relay lenses. A GRIN relay lens generates an intermediate image plane with close to unity magnification. The mounted 0.25pitch objected GRIN lens then images the intermediate image plane and forms a second image plane on the surface of the CMOS imaging sensor. Thanks to Eyal Kimchi at MIT for the initial design of this modification.<br />
<br />
<br clear=all><br />
<br />
{{#ev:youtube|https://www.youtube.com/watch?v=wctf89-i0ZI|640|center}}<br />
<br />
<br clear=all><br />
<br />
[[File:ModScopeOverview.png|center|800px]]<br />
<br />
<br clear=all><br />
<br />
[[File:ModScopePartsOverview.png|center|600px]]<br />
<br />
<br />
<br />
== Miniscope Body Modification ==<br />
[[File:ModScopeHoleMaking.png|right|600px]]<br />
In order to image thin GRIN relay lenses you will need to mount a 0.25pitch objective GRIN lens into the base of your Miniscope. While you could just glue in the lens to the main body of the scope, we prefer using a set screw so the objective lens can be easily removed.<br />
*Step 1: Carefully drill (3/64" bit) a hole on the back side of the main body of the scope. Place the hole as close to the bottom of the scope as possible, making sure to avoid hitting the slot that holds the dichroic mirror. The hole should be aligned with the opening for the GRIN lens.<br />
*Step 2: Tap the newly created hole with a 0-80 tap. This should be done by hand, making sure not to strip the plastic as it is being threaded.<br />
<br />
<br />
<br clear=all><br />
<br />
== GRIN Lens Modification ==<br />
[[File:ModScopeLensSteps.png|right|600px]]<br />
The opening for a GRIN lens in the bottom of the scope is 2.5mm in diameter. Adding a 2.5mm OD protective sleeve around the objective GRIN lens with protect the lens from the set screw as well as align it in the bottom of the scope. K&S 9833 Thin Wall Brass Tube (2.5mm OD x .225mm Wall) works well with 2mm diameter GRIN lenses. This [http://www.mcmaster.com/#8988k32/=111ycx3 precision miniature stainless steel tubing] works well with 1.8mm diameter GRIN lenses.<br />
*Step 1: Use a dremel or jeweler's saw to cut a metal or plastic tube 2mm to 3mm long. File or grind the ends smooth.<br />
*Step 2: Use a drill bit or rod the diameter of the objective GRIN lens to ream out the inside of the tube and clear out any burrs that might have been created during cutting.<br />
*Step 3: Glue or epoxy (do not use super glue due to out gassing) the tube around the objective GRIN lens. We generally apply a thin layer of optical glue around the side of the lens and then slide on the tube. Try to leave only ~0.5mm of exposed lens on the side that will be placed into the scope body. The protective tube can extend all the way to the bottom of the lens or can be left shorter. Care must be taken not to get glue on the top or bottom surface of the lens. Check these surfaces under a stereoscope and clean off any glue with filter paper and ethanol.<br />
<br />
<br clear=all><br />
<br />
= Implant and Imaging =<br />
A nice overview of imaging with a relay GRIN lens + objective GRIN lens can be found in this [http://www.nature.com/nprot/journal/v11/n3/full/nprot.2016.021.html Nature Protocols paper] by Resendez et al. While the approach outlined in this paper is centered around the commercial nVista system from Inscopix, it is directly applicable to our modified Miniscope system with mounted 0.25 pitch objective GRIN lens.<br />
<br />
Our [[Surgery Protocol]] and [https://drive.google.com/file/d/0ByUbjrn9MxK0TWdxUVVjakF3cDQ/view?usp=sharing| Surgery and Baseplating Presentation] focus on how to direct implanting the 0.25pitch objective GRIN lens but the approach outlined in these links is also directly applicable to implanting and imaging thin GRIN relay lenses.</div>Chrisl1026http://miniscope.org/index.php?title=Imaging_With_Thin_GRIN_Lenses&diff=1545Imaging With Thin GRIN Lenses2016-11-14T19:28:01Z<p>Chrisl1026: /* Modification Overview */</p>
<hr />
<div>Want to image with thin GRIN lenses? Follow the simple steps below for modifying your scope to be able to image through 1mm diameter and thinner GRIN relay lenses. Also make sure you have read our [[GRIN Lens Information]] page.<br />
<br />
= Modification Overview =<br />
The Miniscope relies on a 0.25pitch, 2mm or 1.8mm diameter, objective GRIN lens for imaging. This lens can be implanted directly into the brain ([[Surgery Protocol]], [https://drive.google.com/file/d/0ByUbjrn9MxK0TWdxUVVjakF3cDQ/view?usp=sharing| Surgery and Baseplating Presentation]) but it can also be mounted into the scope to allow for imaging through thinner diameter GRIN relay lenses. A GRIN relay lens generates an intermediate image plane with close to unity magnification. The mounted 0.25pitch objected GRIN lens then images the intermediate image plane and forms a second image plane on the surface of the CMOS imaging sensor. Thanks to Eyal Kimchi at MIT for the initial design of this modification.<br />
<br />
[[File:ModScopeOverview.png|center|800px]]<br />
<br />
<br clear=all><br />
<br />
[[File:ModScopePartsOverview.png|center|600px]]<br />
<br />
<br clear=all><br />
<br />
{{#ev:youtube|https://www.youtube.com/watch?v=wctf89-i0ZI|640|center}}<br />
<br />
== Miniscope Body Modification ==<br />
[[File:ModScopeHoleMaking.png|right|600px]]<br />
In order to image thin GRIN relay lenses you will need to mount a 0.25pitch objective GRIN lens into the base of your Miniscope. While you could just glue in the lens to the main body of the scope, we prefer using a set screw so the objective lens can be easily removed.<br />
*Step 1: Carefully drill (3/64" bit) a hole on the back side of the main body of the scope. Place the hole as close to the bottom of the scope as possible, making sure to avoid hitting the slot that holds the dichroic mirror. The hole should be aligned with the opening for the GRIN lens.<br />
*Step 2: Tap the newly created hole with a 0-80 tap. This should be done by hand, making sure not to strip the plastic as it is being threaded.<br />
<br />
<br />
<br clear=all><br />
<br />
== GRIN Lens Modification ==<br />
[[File:ModScopeLensSteps.png|right|600px]]<br />
The opening for a GRIN lens in the bottom of the scope is 2.5mm in diameter. Adding a 2.5mm OD protective sleeve around the objective GRIN lens with protect the lens from the set screw as well as align it in the bottom of the scope. K&S 9833 Thin Wall Brass Tube (2.5mm OD x .225mm Wall) works well with 2mm diameter GRIN lenses. This [http://www.mcmaster.com/#8988k32/=111ycx3 precision miniature stainless steel tubing] works well with 1.8mm diameter GRIN lenses.<br />
*Step 1: Use a dremel or jeweler's saw to cut a metal or plastic tube 2mm to 3mm long. File or grind the ends smooth.<br />
*Step 2: Use a drill bit or rod the diameter of the objective GRIN lens to ream out the inside of the tube and clear out any burrs that might have been created during cutting.<br />
*Step 3: Glue or epoxy (do not use super glue due to out gassing) the tube around the objective GRIN lens. We generally apply a thin layer of optical glue around the side of the lens and then slide on the tube. Try to leave only ~0.5mm of exposed lens on the side that will be placed into the scope body. The protective tube can extend all the way to the bottom of the lens or can be left shorter. Care must be taken not to get glue on the top or bottom surface of the lens. Check these surfaces under a stereoscope and clean off any glue with filter paper and ethanol.<br />
<br />
<br clear=all><br />
<br />
= Implant and Imaging =<br />
A nice overview of imaging with a relay GRIN lens + objective GRIN lens can be found in this [http://www.nature.com/nprot/journal/v11/n3/full/nprot.2016.021.html Nature Protocols paper] by Resendez et al. While the approach outlined in this paper is centered around the commercial nVista system from Inscopix, it is directly applicable to our modified Miniscope system with mounted 0.25 pitch objective GRIN lens.<br />
<br />
Our [[Surgery Protocol]] and [https://drive.google.com/file/d/0ByUbjrn9MxK0TWdxUVVjakF3cDQ/view?usp=sharing| Surgery and Baseplating Presentation] focus on how to direct implanting the 0.25pitch objective GRIN lens but the approach outlined in these links is also directly applicable to implanting and imaging thin GRIN relay lenses.</div>Chrisl1026http://miniscope.org/index.php?title=Imaging_With_Thin_GRIN_Lenses&diff=1544Imaging With Thin GRIN Lenses2016-11-14T19:27:33Z<p>Chrisl1026: /* Modification Overview */</p>
<hr />
<div>Want to image with thin GRIN lenses? Follow the simple steps below for modifying your scope to be able to image through 1mm diameter and thinner GRIN relay lenses. Also make sure you have read our [[GRIN Lens Information]] page.<br />
<br />
= Modification Overview =<br />
The Miniscope relies on a 0.25pitch, 2mm or 1.8mm diameter, objective GRIN lens for imaging. This lens can be implanted directly into the brain ([[Surgery Protocol]], [https://drive.google.com/file/d/0ByUbjrn9MxK0TWdxUVVjakF3cDQ/view?usp=sharing| Surgery and Baseplating Presentation]) but it can also be mounted into the scope to allow for imaging through thinner diameter GRIN relay lenses. A GRIN relay lens generates an intermediate image plane with close to unity magnification. The mounted 0.25pitch objected GRIN lens then images the intermediate image plane and forms a second image plane on the surface of the CMOS imaging sensor. Thanks to Eyal Kimchi at MIT for the initial design of this modification.<br />
<br />
[[File:ModScopeOverview.png|center|800px]]<br />
<br />
<br clear=all><br />
<br />
[[File:ModScopePartsOverview.png|center|600px]]<br />
<br />
{{#ev:youtube|https://www.youtube.com/watch?v=wctf89-i0ZI|640|center}}<br />
<br />
== Miniscope Body Modification ==<br />
[[File:ModScopeHoleMaking.png|right|600px]]<br />
In order to image thin GRIN relay lenses you will need to mount a 0.25pitch objective GRIN lens into the base of your Miniscope. While you could just glue in the lens to the main body of the scope, we prefer using a set screw so the objective lens can be easily removed.<br />
*Step 1: Carefully drill (3/64" bit) a hole on the back side of the main body of the scope. Place the hole as close to the bottom of the scope as possible, making sure to avoid hitting the slot that holds the dichroic mirror. The hole should be aligned with the opening for the GRIN lens.<br />
*Step 2: Tap the newly created hole with a 0-80 tap. This should be done by hand, making sure not to strip the plastic as it is being threaded.<br />
<br />
<br />
<br clear=all><br />
<br />
== GRIN Lens Modification ==<br />
[[File:ModScopeLensSteps.png|right|600px]]<br />
The opening for a GRIN lens in the bottom of the scope is 2.5mm in diameter. Adding a 2.5mm OD protective sleeve around the objective GRIN lens with protect the lens from the set screw as well as align it in the bottom of the scope. K&S 9833 Thin Wall Brass Tube (2.5mm OD x .225mm Wall) works well with 2mm diameter GRIN lenses. This [http://www.mcmaster.com/#8988k32/=111ycx3 precision miniature stainless steel tubing] works well with 1.8mm diameter GRIN lenses.<br />
*Step 1: Use a dremel or jeweler's saw to cut a metal or plastic tube 2mm to 3mm long. File or grind the ends smooth.<br />
*Step 2: Use a drill bit or rod the diameter of the objective GRIN lens to ream out the inside of the tube and clear out any burrs that might have been created during cutting.<br />
*Step 3: Glue or epoxy (do not use super glue due to out gassing) the tube around the objective GRIN lens. We generally apply a thin layer of optical glue around the side of the lens and then slide on the tube. Try to leave only ~0.5mm of exposed lens on the side that will be placed into the scope body. The protective tube can extend all the way to the bottom of the lens or can be left shorter. Care must be taken not to get glue on the top or bottom surface of the lens. Check these surfaces under a stereoscope and clean off any glue with filter paper and ethanol.<br />
<br />
<br clear=all><br />
<br />
= Implant and Imaging =<br />
A nice overview of imaging with a relay GRIN lens + objective GRIN lens can be found in this [http://www.nature.com/nprot/journal/v11/n3/full/nprot.2016.021.html Nature Protocols paper] by Resendez et al. While the approach outlined in this paper is centered around the commercial nVista system from Inscopix, it is directly applicable to our modified Miniscope system with mounted 0.25 pitch objective GRIN lens.<br />
<br />
Our [[Surgery Protocol]] and [https://drive.google.com/file/d/0ByUbjrn9MxK0TWdxUVVjakF3cDQ/view?usp=sharing| Surgery and Baseplating Presentation] focus on how to direct implanting the 0.25pitch objective GRIN lens but the approach outlined in these links is also directly applicable to implanting and imaging thin GRIN relay lenses.</div>Chrisl1026http://miniscope.org/index.php?title=Part_Procurement&diff=1532Part Procurement2016-09-21T22:40:46Z<p>Chrisl1026: /* General Note on PCB Fabrication and Assembly */</p>
<hr />
<div>The miniscope system is consists of a machined plastic housing, custom Printed Circuit Boards (PCBs), and off the shelf components. This guide will take you through the process of ordering everything you need to build a system of your own.<br />
<br />
== General Note on PCB Fabrication and Assembly ==<br />
Pricing for small batch PCB fabrication and assembly is usually dominated by the one time cost of the PCB manufacturer setting up their equipment for the specific order. When ordering 1000's of PCBs this cost is negligible but for orders of 1 to 10 PCBs this cost effectively greatly increases the per board cost. <br />
<br />
We have been contacted by a many labs interested in coordinating larger orders of PCBs to decrease the per board price and have set up a [[Special:WikiForum|forum]] for those labs to get in touch with each other.<br />
<br />
'''For our own labs we usually aim for the following quantities when ordering:'''<br />
;DAQ PCB:<br />
:Fabrication: 20 PCBs. The '''total price''' for fabrication will likely be the same for quantities of 20 or less of this board so you might as well order the maximum number of boards.<br />
:Assembly: 10 PCBs. Assembly is less affected by the one time setup fee so smaller quantities will have a less dramatic per board price change when increasing total quantity. We aim for getting at least 10 PCBs assembled at a time but the cost for doing just 2-4 is still pretty reasonable.<br />
:Price quote you can reference from Sierra Circuits: [http://miniscope.org/images/a/ad/DAQ_PCB_Quote_Sierra_Circuits.pdf DAQ PCB Quote]. The quote for 'Components' is for Sierra Circuits to order all the electrical components for you. The other option would be to order the components yourself and ship them to Sierra Circuits.<br />
;CMOS Imaging Sensor PCB:<br />
:Fabrication: 50 to 100 PCBs. Again, the '''total price''' will likely stay the same for quantities under 50 to 100 of these boards. <br />
:Assembly: 10 PCBs. Just like with the DAQ PCB, per board assembly cost will change less dramatically with smaller quantity orders.<br />
:Price quote you can reference from Sierra Circuits: [http://miniscope.org/images/8/89/CMOS_Imaging_Sensor_PCB_Quote_Sierra_Circuits.pdf CMOS Imaging Sensor PCB Quote]. The quote for 'Components' is for Sierra Circuits to order all the electrical components for you ('''except the CMOS imaging sensor which you can ask to see if they can source as well'''). Please make reference to '''version 3.2''' of the CMOS Imaging sensor PCB when talking with Sierra Circuits and it could be helpful to link to our [https://github.com/daharoni/Miniscope_CMOS_Imaging_Sensor_PCB/tree/master/v3_2 GitHub repository].The other option would be to order the components yourself and ship them to Sierra Circuits.<br />
;Excitation LED PCB<br />
:This board only requires basic fabrication technology which doesn't have a high one time setup fee. We suggest using Silver Circuits PCB Fabricators for this board.<br />
<br />
== Miniscope Master Parts List ==<br />
An up-to-date list of all components (besides electrical components listed in the each PCB project's Bill of Materials (BoM)) can be found in our Google Documents [https://docs.google.com/spreadsheets/d/12H71DU2QX8d7efUE4yNuikBEiIzKaXjYqdc0A-oLNSw/edit?usp=sharing parts list]. All components on this list should have less than a two week lead time except<br />
*GRIN lenses (see [[GRIN Lens Information]])<br />
*CMOS Imaging Sensor<br />
After running through the procurement details below make sure to double check with the parts list that all components have been ordered.<br />
== Head Mounted Scope ==<br />
=== Machined Housing Parts ===<br />
The design files for all parts that need to be machined can be found on the [https://github.com/daharoni/Miniscope_Machined_Parts.git Miniscope Machined Parts Github Repository]. We have included both the Solidworks files as well as exported each part as a .stl.<br />
<br />
The three housing components (MainBody, FocusingSlider, FilterSetCover) are CNC machined out of Delrin plastic. While the parts themselves are not too complex, their small size and thin walls makes them tricky to machine. Your campus machine shop should be able to machine these parts for you but generally this will take longer and be significantly more expensive than sending the parts out for machining.<br />
<br />
Shylo Stiteler is a local machinist that is very familiar with our project. You can contact Shylo by email, shylostiteler@gmail.com, with the parts you need machined. His work is very reasonably priced, quick, and consistent.<br />
<br />
We also should mention [https://www.protolabs.com/ Protolabs] for general machining projects. They are very quick and reliable but will not machine parts with wall thickness under 1mm. This means they won't be able to machine the majority of scope parts without added addition mass.<br />
<br />
=== Machined Aluminum Baseplate ===<br />
The design file for the Baseplate can also be found on the [https://github.com/daharoni/Miniscope_Machined_Parts.git Miniscope Machined Parts Github Repository]. Similar to what is stated above, this part can likely be machined by your local machine shop but we recommend getting the part machined by Shylo Stiteler's company, shylostiteler@gmail.com.<br />
<br />
=== Optical Components ===<br />
The optical components in the microscope are all off the shelf components which generally have less than a week lead time to arrive.<br />
;Diced filter set<br />
:We use Chroma filters sets in all our scopes but have also had success with Semrock and Omega Optical filters. All three of these companies will dice their standard filters to what ever size you need. The filter set and dice dimensions we use can be found on our [https://docs.google.com/spreadsheets/d/12H71DU2QX8d7efUE4yNuikBEiIzKaXjYqdc0A-oLNSw/edit?usp=sharing parts list], and you can contact Dick Stewart <dstewart@chroma.com> for a price quote.<br />
;Internal lenses<br />
:Both the half-ball lens and achromatic lens can be purchased through [http://www.edmundoptics.com/ Edmund Optics]. Part numbers can be found on our [https://docs.google.com/spreadsheets/d/12H71DU2QX8d7efUE4yNuikBEiIzKaXjYqdc0A-oLNSw/edit?usp=sharing parts list].<br />
;GRIN lenses<br />
:Acquiring GRIN lenses may prove a bit more complicated than acquiring the rest of the components in our system. For this reason we have a [[GRIN Lens Information|dedicated GRIN lens page]] on this topic.<br />
<br />
=== CMOS Imaging Sensor PCB ===<br />
'''IMPORTANT''': If you downloaded the CMOS Imaging Sensor PCB files before January 27, 2015 please update them with the current version on GitHub.<br />
<br />
Design, fabrication, and assembly files can be found that the [https://github.com/daharoni/Miniscope_CMOS_Imaging_Sensor_PCB.git Miniscope CMOS imaging sensor PCB Github repository]. The newest version of the PCB can be found in folder 'v3_2'.<br />
<br />
The 'Fabrication Files' folder contains all the information needed to get the PCB fabricated. PCB fabrication means the PCB is printed but not assembled. The PCB is a 4 layer, 0.031" thick FR4 circuit board with In Pad Vias.<br />
*'CMOS_v3_2_PCB_Fab_Info.docx' has all the relevant specs for PCB fabrication.<br />
<br />
The 'Assembly Files' filder contains all the information needed to get the fabricated circuit board assembled with electrical components.<br />
*'Serializer_System2_MT9V032_BOM.xlsx' lists all electrical components needed for assembly along with each components PCB designator. A PCB assembly house can likely offer a turn-key assembly solution where they will hand the ordering of all these components for a small markup. You can also order them yourself and ship them as a kit to the assembler.<br />
*'CMOS_v3_2_Assy_Info.docx' has all the relevant information for PCB assembly.<br />
<br />
While it is possible to assemble this PCB yourself, we suggest most labs send it out to a professional assembly house. For both fabrication and assembly we recommend using [https://www.protoexpress.com/ Sierra Circuits]. The gerber files and board information document are needed for fabrication. The BOM (Bill of Materials) file and PnP (Pick and Place) file are needed for assembly. Generally PCB assembly houses can provide turnkey services which means they will not only assembly the PCB but also handle ordering all the individual components for a small markup. <br />
<br />
In order to minimize the size of this PCB, buried vias and two sided assembly are used which results in a higher production costs than more standard boards. When getting a price quote we suggest asking for pricing at a few different quantities (5, 10, 20 boards). At these small quantities there is usually a range of quantities where the overall price remains practically the same for fabricating and assembly more boards.<br />
<br />
:''' IMPORTANT: Ordering the Imaging Sensor:''' All electrical components can be easily purchased through Digikey except for the imaging sensor. You should order the imaging sensor yourself and send it in to be assembled ('''Note:''' Currently Sierra Circuits should be able to source the imaging sensor as well so ask when requesting a quote). The CMOS imaging sensor used is the Aptina (or OnSemiconductor) MT9V032C12STM and can be found through Arrow Electronics or Aliexpress.<br />
<br />
=== Excitation LED PCB ===<br />
Design and fabrication files can be found at the [https://github.com/daharoni/Miniscope_Excitation_LED_PCB.git Miniscope Excitation LED PCB GitHub Repository]. <br />
<br />
The technology needed to fabrication this board is much simpler than what is needed for the CMOS imaging sensor PCB and DAQ PCB. For this reason we suggest using the 'PCB Production' option through [[http://www.custompcb.com/ Silver Circuits]]. You can upload the gerber files directly to their site for a price quote but we suggest emailing them to get a better deal.<br />
<br />
If you want to get the standard Excitation LED PCBs made you can just email sales@custompcb.com and tell them that you would like to purchase Order # 7500 (no need to include any design/fabrication files). This will give you 125 PCB for a little over $200. While 125 PCBs (the minimum order) is more than what is needed for most labs, the total price will still be significantly cheaper than most other PCB fabrication companies.<br />
<br />
'''Fabrication Properties:'''<br />
*Number of layers: 2<br />
*Substrate: 0.031” FR4<br />
*Board width: 5mm<br />
*Board height: 7.5mm<br />
*Copper weight: 1oz (not important)<br />
*Surface: HASL or ENIG (not important)<br />
*Board shape: Custom<br />
*Cutouts: None<br />
*Solder mask color: Black<br />
*Sinkscreen layers: Bottom<br />
*Sinkscreen color: White<br />
*Electrical Test: Not required<br />
*Routing: Either individual or panalized with v-scores<br />
*Ask to get them panelized on 4"x5" PCBs<br />
<br />
=== Miscellaneous Parts ===<br />
<br />
== Data Acquisition System ==<br />
The DAQ system is composed of a DAQ PCB and 3D printed housing. The DAQ PCB needs to be fabricated and assembled by a PCB Fab and Assembly House capable of handling prototype and low volume orders (we suggest Sierra Circuits). In general you will mainly be paying for the setup cost when getting low volumes of PCBs fabricated and assembled. For this reason it is cost effective to ask for a price quote at multiple quantities (1, 5, 10, 20) to get an idea of price breakdown. For these DAQ PCBs it will cost around the same amount to get 1, 5, 10 produced.<br />
<br />
=== Data Acquisition PCB ===<br />
Design, fabrication, and assembly files can be found at the [https://github.com/daharoni/Miniscope_DAQ_PCB.git Miniscope DAQ PCB Github repository].<br />
<br />
The 'Fabrication Files' folder contains all the information needed to get the PCB fabricated. PCB fabrication means the PCB is printed but not assembled. The PCB is a 4 layer, 0.062" thick FR4 circuit board with controlled impedance and tented vias. The DAQ board contains a 0.8mm pitch BGA component so the surface finish needs to be ENIG. <br />
*'Miniscope_DAQ_PCB_Fab_Info.docx' has all the relevant specs for PCB fabrication.<br />
<br />
The 'Assembly Files' filder contains all the information needed to get the fabricated circuit board assembled with electrical components. I usually have all components except the through-hole components assembly by an Assembly House but you can have the through-hole components assembled by them as well. You will need to modify the Bill of Materials (BOM) depending on which components you want assembled.<br />
*'USB_Control_FPD_Linkv2_BOM.xlsx' lists all electrical components needed for assembly along with each components PCB designator. A PCB assembly house can likely offer a turn-key assembly solution where they will hand the ordering of all these components for a small markup. You can also order them yourself and ship them as a kit to the assembler.<br />
*'Miniscope_DAQ_PCB_Assy_Info.docx' has all the relevant information for PCB assembly.<br />
<br />
=== DAQ PCB Case ===<br />
We have designed a simple case to enclose and protect the DAQ PCB which can be 3D printed [http://shpws.me/Kf3H here].<br />
<br />
=== Miscellaneous Parts ===<br />
<br />
== Cabling and Connector==<br />
On the 'Head Mounted Scope' page of the [https://docs.google.com/spreadsheets/d/12H71DU2QX8d7efUE4yNuikBEiIzKaXjYqdc0A-oLNSw/edit?usp=sharing parts list] you will find part numbers for 2 coaxial cable options as well as the suggested SMA coaxial connector. Even if you are experienced with soldering, we suggest using the [https://github.com/daharoni/Miniscope_Coax_2_SMA_PCB Coax2SMA PCB] along with an edge mounted SMA connector to connectorize the coax cables. The Coax2SMA PCB is a very simple PCB that any PCB fabrication house can produce. We suggest using the link below to have these PCBs fabricated by OSH Park (very quick, cheap, and easy).<br />
<br />
[https://oshpark.com/shared_projects/xtQGQ32E Order from OSH Park]<br />
<br />
== Tools ==<br />
There is a dedicated page of the [https://docs.google.com/spreadsheets/d/12H71DU2QX8d7efUE4yNuikBEiIzKaXjYqdc0A-oLNSw/edit?usp=sharing parts list] outlining the tools you will need for assembly of the Miniscope system.</div>Chrisl1026http://miniscope.org/index.php?title=Initial_Testing_of_Assembled_Miniscopes&diff=1531Initial Testing of Assembled Miniscopes2016-09-21T22:12:29Z<p>Chrisl1026: /* Imaging GFP Slides */</p>
<hr />
<div>'''This page is a work in progress.'''<br />
<br />
Once your Miniscope system is up and running, it is important to be able to test, and debug, all aspects of the system before moving to imaging in vivo. The sections below will discuss the procedures we use to validate Miniscopes we build before using them in experiments. As you become more comfortable with using Miniscopes some of the sections below can be skipped.<br />
<br />
== Testing the Coaxial Cable Connection ==<br />
In our experience, the connection of the coax cable, either to the CMOS Imaging Sensor PCB or SMA connector, is by far the most common point of failure when building a Miniscope System. Take care when soldering these connections not to short the inner conductor to the outer shield. It helps to cover the solder joint as well as ~1cm of coax cable extending from the solder join in a semi-flexible epoxy, silicone, or glue (hot glue works well) to take the strain of cable movement off of the solder joint.<br />
<br />
Once assembled, connect the Miniscope system to your computer and run the DAQ software. With the video streaming from the scope, move/twist/wiggle the coax cable with greater intensity that what you would expect an animal to apply, especially at the ends of the cable. If soldered correctly, the video stream should not drop out even with excessive movement and twisting of the cable.<br />
<br />
== Testing the Stability of the DAQ Software ==<br />
[[File:SoftwareTests.png|right|500px]]<br />
When running your Miniscope system on a new computer it is important to check the following<br />
*'''Stability of video stream:''' We have found some combinations of USB drivers, computer hardware, and Window's OS can lead to the video stream failing a few minutes after the software has connected to the scope. This seems to mainly be an issue with Windows 8 and is independent of if you are recording the video to disk. To test the stability of your system, <br />
*#Connect a scope to the DAQ Box and then the DAQ Box to the computer.<br />
*#Open up the DAQ software and connect to the Miniscope.<br />
*#Leave the system running for 5 minutes. Do not click the 'record' button.<br />
*#If the video stream is still present (not Red Screen of Doom) your system should not have any driver or OS issues.<br />
*'''Streaming video frame rate:''' The default frame rate of your Miniscope system is 30FPS but can be adjusted using the drop down 'Frame Rate' box in our DAQ software (labeled with a red '1' in the picture to the right). <br />
*#Connect a scope to the DAQ Box and then the DAQ Box to the computer.<br />
*#Open up the DAQ software and connect to the Miniscope.<br />
*#Observe the current frame rate (labeled with a red '2' in the picture to the right). It should be stable within 1 FPS from the expected value. This is just an approximate measure of the frame rate, the frame rate of the recorded video should be extremely stable.<br />
*#You can also connect to a behavioral camera to add strain on your system. Generally the behavioral camera will have larger fluctuations in the displayed frame rate.<br />
*'''Write speed of video data:''' You want to make sure your computer is able to write the uncompressed video data to your HDD or SSD as quick (and hopefully much quicker)than the rate at which you are acquiring it. Slow or encryted hard drives can be a source of problems here.<br />
*#Connect a scope to the DAQ Box and then the DAQ Box to the computer.<br />
*#Open up the DAQ software and connect to the Miniscope.<br />
*#Click 'record' and observe the 'Write Speed (fps)' (labled with a red '3' in the picture to the right). This box displays the current write speed of your data. The number displayed will fluctuate but should stay above the acquisition frame rate of your Miniscope. If the write speed falls below the acquisition frame rate, video frames will be written into a circular buffer which is 256 frames long. If the write speed stays low for too long the software will begin to overwrite frames in the circular buffer. The 'timestamps.dat' file created during recording keeps track of the buffer size and is an addition place you can look when evaluating the write speed of your computer.<br />
<br clear=all><br />
<br />
== Imaging Your Surroundings ==<br />
[[File:ImagingWithoutGRINLens.png|right|500px]]<br />
An assembled scope can image your surrounds when a GRIN lens is '''not placed''' into the hole in the base of the scope. Connect the scope to your computer and then point the base of the scope toward objects that are illuminated with room or sun light.<br />
*The scope has a green bandpass filter sitting before the CMOS imaging sensor. This means you will only be collecting light between 500nm and 550nm.<br />
*Most objects in the environment will not fluoresce under the blue excitation LED. This means most object will not show up unless being illuminated by a light source that contains green wavelengths such as room light or sunlight. <br />
*Adjust the focus slider to adjust the focal plane of your Miniscope. Most object will appear to be "infinitely" far away from the CMOS imaging sensor. Your focus slider will need to be placed close to its highest point in order to focus on these objects. This focusing slider position is equivalent to imaging at the bottom surface of your GRIN lens if a GRIN lens was mounted into the base of the scope.<br />
<br />
<br clear=all><br />
<br />
== Checking for Light Leakage ==<br />
[[File:LightLeakage.png|right|500px]]<br />
When assembled correctly, no excitation light from the LED should leak onto the CMOS imaging sensor in your scope. The following steps will walk you through testing a scope for such light leakage. If you do find that your scope leaks light the most common sources of the leak are the excitation or emission filter being scratched, placed in the wrong orientation, or significantly misaligned.<br />
#Connect a scope to the DAQ Box and then the DAQ Box to the computer.<br />
#Open up the DAQ software and connect to the Miniscope.<br />
#Place the opening in the base of the scope (you can do this with or without a GRIN lens) against a black surface that won't fluoresce. You may be surprised at what materials weakly fluoresce green when blasted with blue excitation light.<br />
#Turn the exposure and gain of the scope to their maximum values.<br />
#Slowly turn up the excitation LED power from 0% to max power. Depending on your scope and DAQ software version the LED will likely max out at around 40%.<br />
#As you increase the LED power watch the video stream for large increases in pixel brightness across large regions of your image. A small overall increase in pixel value of ~20 (the pixel values range from 0 to 255) is expected. You may also notice at max gain that some pixels become noisy. This is also normal but care should be taken in experiments to minimize this noise by limiting the gain or by correct these noisy pixels during offline processing of your data.<br />
<br />
<br clear=all><br />
<br />
== Imaging Calibration Slides ==<br />
[[File:ImagingTestSlide.png|right|500px]]<br />
[[File:MiniscopeCalibrationSlide.png|thumb|right|500px|Image of a calibration slide with 9.8μm line spacing (superimposed red boxes are 10px x 10px)]]<br />
<br />
This is probably the most important initial test you can do with your Miniscope system. Not only can it uncover issues in your system but will also give you a good sense of how the optics, imaging, and focusing slider work. The figure to the right shows a modified test/calibration slide (left), optional GRIN lens holder (middle), and example configuration for imaging a test slide (right).<br />
*The modified test/calibration slide is a [http://www.thorlabs.us/navigation.cfm?guide_id=2332 resolution test slide] with [http://www.amazon.com/JVCC-Stage-Set-Spike-Tape-Fluorescent/dp/B000QDVNH0/ref=sr_1_12?s=industrial&srs=2529683011&ie=UTF8&qid=1461180753&sr=1-12 green fluorescing tape] attached to the underside of the glass. It is important that the printed surface of the test slide be the side that is closest to the GRIN lens. Different thicknesses of cover glass can be used between the test slide and GRIN lens to get a feeling for imaging at different depths.<br />
*The middle picture shows a simple GRIN lens holder we had made. This holder is a block of Delrin plastic with different diameter holes drilled into it. a GRIN lens can be placed into the holder and then set on top of the modified test slide. A Miniscope can then be set onto of the holder to image the slide. The holder does a nice job keeping the surface of the GRIN lens co-planer to the surface of the test slide.<br />
*The right picture should an example configuration to image a test slide. This picture is of a scope that has been modified, [Imaging With Thin GRIN Lenses], to hold a GRIN lens in its base. While the picture shows the scope being held by hand, one could easily mount the scope in a clamp for more stable imaging. If using a GRIN lens holder as described above, a similar configuration would be used except the GRIN lens and holder would be in place of the bare GRIN lens.<br />
<br />
The image on the bottom right is of a calibration slide with 9.8μm line spacing (superimposed red boxes are 10px x 10px).<br />
<br clear=all><br />
<br />
== Imaging GFP Slides ==<br />
{{#ev:youtube|https://youtu.be/lxEUkP-YI8g|640|right}}<br />
Similar to imaging test slides, you can use your Miniscope to image brain slice slides expressing GFP or slides with other fluorescent sources. A few key points for imaging slides are listed below:<br />
*Your scope has a green bandpass filter (500nm to 550nm) in front the the CMOS imaging sensor. This means only will only be able to see green fluorescence.<br />
*Take note of the thickness of your cover slip, usually around 170um thick. You will be imaging through the cover slip to reach the fluorescing sample. If you are using less than a 15mm focal length achromatic lens, you will likely not be able to focus past the bottom of the cover slip. Even a 15mm focal length achromatic lens will generally just reach about 50um below the bottom of the cover slip with the focusing slider pushed to its lowest position. <br />
<br />
The video to the right is of a TetTag GFP slide being imaged with a 15mm focal length achromatic lens and 0.25 pitch GRIN lens. A few notes:<br />
*Their are some compression artifacts present due to YouTube's compression of the video. <br />
*The disk of light near the center of the field of view is due to reflection of the fluorescent light between the cover slip and bottom of GRIN lens... you should not see this disk when imaging in vivo. <br />
*The left and right portions of the field of view are dimmer than what you should expect to see with your own system. The outer dimness was greatly reduced in the newer versions of the Miniscope main body.</div>Chrisl1026http://miniscope.org/index.php?title=Imaging_With_Thin_GRIN_Lenses&diff=1530Imaging With Thin GRIN Lenses2016-09-21T22:05:50Z<p>Chrisl1026: /* GRIN Lens Modification */</p>
<hr />
<div>Want to image with thin GRIN lenses? Follow the simple steps below for modifying your scope to be able to image through 1mm diameter and thinner GRIN relay lenses. Also make sure you have read our [[GRIN Lens Information]] page.<br />
<br />
= Modification Overview =<br />
The Miniscope relies on a 0.25pitch, 2mm or 1.8mm diameter, objective GRIN lens for imaging. This lens can be implanted directly into the brain ([[Surgery Protocol]], [https://drive.google.com/file/d/0ByUbjrn9MxK0TWdxUVVjakF3cDQ/view?usp=sharing| Surgery and Baseplating Presentation]) but it can also be mounted into the scope to allow for imaging through thinner diameter GRIN relay lenses. A GRIN relay lens generates an intermediate image plane with close to unity magnification. The mounted 0.25pitch objected GRIN lens then images the intermediate image plane and forms a second image plane on the surface of the CMOS imaging sensor. Thanks to Eyal Kimchi at MIT for the initial design of this modification.<br />
<br />
[[File:ModScopeOverview.png|center|800px]]<br />
<br />
<br clear=all><br />
<br />
[[File:ModScopePartsOverview.png|center|600px]]<br />
<br />
== Miniscope Body Modification ==<br />
[[File:ModScopeHoleMaking.png|right|600px]]<br />
In order to image thin GRIN relay lenses you will need to mount a 0.25pitch objective GRIN lens into the base of your Miniscope. While you could just glue in the lens to the main body of the scope, we prefer using a set screw so the objective lens can be easily removed.<br />
*Step 1: Carefully drill (3/64" bit) a hole on the back side of the main body of the scope. Place the hole as close to the bottom of the scope as possible, making sure to avoid hitting the slot that holds the dichroic mirror. The hole should be aligned with the opening for the GRIN lens.<br />
*Step 2: Tap the newly created hole with a 0-80 tap. This should be done by hand, making sure not to strip the plastic as it is being threaded.<br />
<br />
<br />
<br clear=all><br />
<br />
== GRIN Lens Modification ==<br />
[[File:ModScopeLensSteps.png|right|600px]]<br />
The opening for a GRIN lens in the bottom of the scope is 2.5mm in diameter. Adding a 2.5mm OD protective sleeve around the objective GRIN lens with protect the lens from the set screw as well as align it in the bottom of the scope. K&S 9833 Thin Wall Brass Tube (2.5mm OD x .225mm Wall) works well with 2mm diameter GRIN lenses. This [http://www.mcmaster.com/#8988k32/=111ycx3 precision miniature stainless steel tubing] works well with 1.8mm diameter GRIN lenses.<br />
*Step 1: Use a dremel or jeweler's saw to cut a metal or plastic tube 2mm to 3mm long. File or grind the ends smooth.<br />
*Step 2: Use a drill bit or rod the diameter of the objective GRIN lens to ream out the inside of the tube and clear out any burrs that might have been created during cutting.<br />
*Step 3: Glue or epoxy (do not use super glue due to out gassing) the tube around the objective GRIN lens. We generally apply a thin layer of optical glue around the side of the lens and then slide on the tube. Try to leave only ~0.5mm of exposed lens on the side that will be placed into the scope body. The protective tube can extend all the way to the bottom of the lens or can be left shorter. Care must be taken not to get glue on the top or bottom surface of the lens. Check these surfaces under a stereoscope and clean off any glue with filter paper and ethanol.<br />
<br />
<br clear=all><br />
<br />
= Implant and Imaging =<br />
A nice overview of imaging with a relay GRIN lens + objective GRIN lens can be found in this [http://www.nature.com/nprot/journal/v11/n3/full/nprot.2016.021.html Nature Protocols paper] by Resendez et al. While the approach outlined in this paper is centered around the commercial nVista system from Inscopix, it is directly applicable to our modified Miniscope system with mounted 0.25 pitch objective GRIN lens.<br />
<br />
Our [[Surgery Protocol]] and [https://drive.google.com/file/d/0ByUbjrn9MxK0TWdxUVVjakF3cDQ/view?usp=sharing| Surgery and Baseplating Presentation] focus on how to direct implanting the 0.25pitch objective GRIN lens but the approach outlined in these links is also directly applicable to implanting and imaging thin GRIN relay lenses.</div>Chrisl1026http://miniscope.org/index.php?title=Imaging_With_Thin_GRIN_Lenses&diff=1529Imaging With Thin GRIN Lenses2016-09-21T22:04:29Z<p>Chrisl1026: /* Miniscope Body Modification */</p>
<hr />
<div>Want to image with thin GRIN lenses? Follow the simple steps below for modifying your scope to be able to image through 1mm diameter and thinner GRIN relay lenses. Also make sure you have read our [[GRIN Lens Information]] page.<br />
<br />
= Modification Overview =<br />
The Miniscope relies on a 0.25pitch, 2mm or 1.8mm diameter, objective GRIN lens for imaging. This lens can be implanted directly into the brain ([[Surgery Protocol]], [https://drive.google.com/file/d/0ByUbjrn9MxK0TWdxUVVjakF3cDQ/view?usp=sharing| Surgery and Baseplating Presentation]) but it can also be mounted into the scope to allow for imaging through thinner diameter GRIN relay lenses. A GRIN relay lens generates an intermediate image plane with close to unity magnification. The mounted 0.25pitch objected GRIN lens then images the intermediate image plane and forms a second image plane on the surface of the CMOS imaging sensor. Thanks to Eyal Kimchi at MIT for the initial design of this modification.<br />
<br />
[[File:ModScopeOverview.png|center|800px]]<br />
<br />
<br clear=all><br />
<br />
[[File:ModScopePartsOverview.png|center|600px]]<br />
<br />
== Miniscope Body Modification ==<br />
[[File:ModScopeHoleMaking.png|right|600px]]<br />
In order to image thin GRIN relay lenses you will need to mount a 0.25pitch objective GRIN lens into the base of your Miniscope. While you could just glue in the lens to the main body of the scope, we prefer using a set screw so the objective lens can be easily removed.<br />
*Step 1: Carefully drill (3/64" bit) a hole on the back side of the main body of the scope. Place the hole as close to the bottom of the scope as possible, making sure to avoid hitting the slot that holds the dichroic mirror. The hole should be aligned with the opening for the GRIN lens.<br />
*Step 2: Tap the newly created hole with a 0-80 tap. This should be done by hand, making sure not to strip the plastic as it is being threaded.<br />
<br />
<br />
<br clear=all><br />
<br />
== GRIN Lens Modification ==<br />
[[File:ModScopeLensSteps.png|right|600px]]<br />
The opening for a GRIN lens in the bottom of the scope is 2.5mm in diameter. Adding a 2.5mm OD protective sleeve around the objective GRIN lens with protect the lens from the set screw as well as align it in the bottom of the scope. K&S 9833 Thin Wall Brass Tube (2.5mm OD x .225mm Wall) works well with 2mm diameter GRIN lenses. This [http://www.mcmaster.com/#8988k32/=111ycx3 precision miniature stainless steal tubing] works well with 1.8mm diameter GRIN lenses.<br />
*Step 1: Use a dremel or jeweler's saw to cut a metal or plastic tube 2mm to 3mm long. File or grind the ends smooth.<br />
*Step 2: Use a drill bit or rod the diameter of the objective GRIN lens to ream out the inside of the tube and clear out any burrs that might have been created during cutting.<br />
*Step 3: Glue or epoxy (do not use super glue due to out gassing) the tube around the objective GRIN lens. We generally apply a thin layer of optical glue around the side of the lens and then slide on the tube. Try to leave only ~0.5mm of exposed lens on the side that will be placed into the scope body. The protective tube can extend all the way to the bottom of the lens or can be left shorter. Care must be taken not to get glue on the top or bottom surface of the lens. Check these surfaces under a stereoscope and clean off any glue with filter paper and ethanol.<br />
<br />
<br clear=all><br />
<br />
= Implant and Imaging =<br />
A nice overview of imaging with a relay GRIN lens + objective GRIN lens can be found in this [http://www.nature.com/nprot/journal/v11/n3/full/nprot.2016.021.html Nature Protocols paper] by Resendez et al. While the approach outlined in this paper is centered around the commercial nVista system from Inscopix, it is directly applicable to our modified Miniscope system with mounted 0.25 pitch objective GRIN lens.<br />
<br />
Our [[Surgery Protocol]] and [https://drive.google.com/file/d/0ByUbjrn9MxK0TWdxUVVjakF3cDQ/view?usp=sharing| Surgery and Baseplating Presentation] focus on how to direct implanting the 0.25pitch objective GRIN lens but the approach outlined in these links is also directly applicable to implanting and imaging thin GRIN relay lenses.</div>Chrisl1026http://miniscope.org/index.php?title=Animal_Behavior_Guide&diff=1528Animal Behavior Guide2016-09-21T21:59:57Z<p>Chrisl1026: /* Starting an experiment */</p>
<hr />
<div>Here are some tips to for using the miniscopes to run experiments in freely moving mice.<br />
<br />
<br />
== Experimental timeline ==<br />
Plan you experiments schedule accordingly. Here is a typical timeline for getting mice ready for experiments in freely moving mice.<br />
<br />
:Week 1: virus injection surgery<br />
:Week 2: GRIN lens implant surgery<br />
:Week 3: recovery<br />
:Week 4: base plate surgery + acclimation to experimenters/handling<br />
:Week 5: habituation to wearing miniscopes & setting imaging focal plane<br />
:Week 6: begin experiment!<br />
<br />
== Acclimation to experimenters/Handling ==<br />
We typically wait at least a day after base plate surgery to begin handling. We recommend that mice become well acclimated to the experimenters prior to mounting miniscopes on awake moving mice. It is important that the mice feel comfortable with the experimenters as stress during the mounting of the miniscope during the experiment can cause adverse effects for the mice and thus, experiment. After gently handling the animals a few times, acclimate the mice to getting something placed on and off their heads by gently taking the protective caps on and off their base plate (that is cemented on the mice's heads). The experimenters should be able to quickly remove the cap without much struggle from the animal before continuing onto the habituation phase.<br />
<br />
<br />
== Habituation to wearing miniscopes ==<br />
Once the mice (and experimenters) are comfortable with the cap removal, habituate the mice to wearing the miniscopes while freely moving. To habituate, one experimenter should calmly hold the mouse and remove the cap, while the second experimenter replaces the cap with the miniscope. The miniscope should easily snap into place with the magnets. The second experimenter should gently hold the sides of the base plate with one hand while securing the miniscope with the set screw with the other hand. Especially during the first time, the animal may struggle. If the animal is struggling too much, let go and calm the animal. If the animal resists and struggles TOO much while you are holding onto the base plate, it can lead to the dental cement holding the base plate to come off the skull. When the animal is calm, try to hold the base plate and insert the set screw to secure the miniscope again. Once the miniscope is secured, let the animal explore your habituation environment of choice. We typically just let them move freely around in their home cage (even if there are other mice in the cage). Depending on your experimental demands, you will need to habituate to different levels of movement. If the experiment only requires mice to walk around and explore, we recommend habituating for at least 3 days for 10 minutes each day. If the experiment requires the mice to run swiftly (e.g. down a linear track), then we recommend habituating for at least 5 days for 10-30 minutes each day.<br />
<br />
== Setting imaging focal plane & LED intensity ==<br />
We recommend setting the focal plane of imaging and LED intensity prior to the experimental day. This can be done during the habituation sessions. Move the focus slider along the neck of the body of the miniscope until you've reached your optimal focal plane of the brain. To set the focal plane, tighten the set screw into the focus slider until the focus slider is secure. Be careful of over-tightening, as the screw can make indents into to the plastic material of the neck of the body. Once the focal plane is set, do not change it for the duration of your experiment if you need to image the same set of cells across sessions. Typically, we have found that for some mice, they can use the same miniscope set at a certain focal plane. For other mice, they need the miniscope to be set to a different height.<br />
<br />
During this session, you may also want to find the optimal level of LED intensity for imaging. Make sure that the there is enough blue light to see both the firing of the cells as well as some blood vessels to be used as landmarks to align frames. We recommend taking a video during the habituation session with your settings and analyzing the video to make sure the settings are optimal prior to the start of the experiment. Also, take a snapshot of the brain that you will use to reference for all future recording sessions. That way, you can begin your experiment with confidence and ease!<br />
<br />
== Starting an experiment ==<br />
Here is an example checklist that can be used.<br />
<br />
*Check if there is enough space on the hard drive for the data files that will be collected throughout the experiment. If you are using a behavioral camera, don't forget to include that as well.<br />
*Check and clean the imaging sensor before screwing it on the focus slider. If there is dust or dirt, use lens paper and alcohol to gently clean the sensor. This typically only needs to be done once prior to securing onto the focus slide. If the imaging sensor is being removed between sessions, make sure to screw it back onto the focus slide in the same orientation!<br />
*Connect the miniscope cable with the DAQ, connect the USB 3.0 cable with the DAQ and computer, and open the Miniscope Control application on the computer.<br />
*Make sure the 3 lights on the DAQ turn on and the red light on the miniscope turns on.<br />
*Make sure that the miniscope is flush against the base plate or as much as possible and the miniscope is not tilted and the set screw is not in the way.<br />
*Compare the live image you see with the snapshot of the same region you took during the habituation. Try to match it up as best as possible. The blood vessels, contrast, brightness should all look similar. If it doesn't then troubleshoot as to why it is different.<br />
*Gently tighten the set screw to secure the miniscope onto the base plate. Stop once the screw makes contact with the plastic of the miniscope. Over tightening will make an indentation into the plastic body and cause damage. To check if the miniscope is secure, GENTLY wiggle and tug on the miniscope while attached to the base plate while watching the imaging of the the brain. Moving the miniscope should not cause movement of the image on the video.<br />
*Make sure to have enough slack on the cable so the mice can move freely but not so much that the mice can chew on the cable. If you're not using a commutator, attend to the possible twisting of the cables while the animals are moving.<br />
*Check to see that the write speed of the computer exceeds the FPS collected. <br />
*After the trial, visually inspect that data folder to make sure the data was correctly collected.<br />
*Back up your data!!!</div>Chrisl1026http://miniscope.org/index.php?title=Surgery_Protocol&diff=1527Surgery Protocol2016-09-21T04:15:15Z<p>Chrisl1026: /* Basic Equipment needed */</p>
<hr />
<div>The following is a basic outline of our procedure for implanting a GRIN lens above the hippocampus for CA1 imaging. For videos of surgery and baseplating, see the [[Online Workshop]]. Also, see the recent Nature Protocols paper from the Stuber Lab ([http://www.nature.com/nprot/journal/v11/n3/full/nprot.2016.021.html link]) that has a very detailed description of the surgery. <br />
<br />
In order to image, animals will need to undergo three surgical procedures: a virus injection, a GRIN lens implantation, and baseplate attachment.<br />
<br />
== Virus Injection ==<br />
Before implanting the GRIN lens, you will need to inject a fluorescent indicator such as GCaMP6. We have generally used AAV1.Syn.GCaMP6f.WPRE.SV40 from Penn Vector (Item number AV-1-PV2822) and had great success in dorsal CA1. You may also be able to use GCaMP6 transgenic mice, but we have not tested whether any of the newer lines are bright enough to be imaged.<br />
<br />
== GRIN Lens Implantation==<br />
<br />
=== Basic Equipment Needed ===<br />
You will only need basic surgical equipment to perform GRIN lens implantation surgeries. This includes a mouse stereotax, an isoflurane vaporizor, surgical heat pad, stereo surgical microscope, light source, dental drill, and bead sterilizer. For recommended equipment, see the master [https://docs.google.com/spreadsheets/d/12H71DU2QX8d7efUE4yNuikBEiIzKaXjYqdc0A-oLNSw/edit?usp=sharing parts list].<br />
<br />
You will need the following basic tools and supplies: fine forceps, blunt forceps, fine scissors, scalpel and blade, small skull screws, drill bits, cyanoacrylate glue, dental cement, Kwik-Sil, and [[cortex buffer]]<br />
<br />
=== Aspirator and Lens Holder ===<br />
You will need an aspirator to remove cortex above the hippocampus. This can be built very simply with a vacuum line (or pump), a liquid trap, 1 ml syringe (with a hole to control suction), blunt needles, tubing and connectors, and hot glue.<br />
[[File:Aspirator.PNG|center|500px]]<br />
<br />
You can easily build a GRIN lens holder by using a suction system to hold the lens in place as it is dropped into the brain for implantation. To build, start with two 1ml micropipette tips. Cut the first so that the tip is just larger than the diameter of the GRIN lens you will be using. Cut the second tip so that the end is just smaller than the size of the GRIN lens. Connect this end to a 1ml syringe and connect to a vacuum line. This will then hold the lens in place during implantation. You can then use tape to attach this holder to a stereotax. Another solution we have used is a basic drill chuck, but this is only good for large (2mm) lenses.<br />
[[File:LensHolder.PNG|center|500px]]<br />
<br />
=== Skull Preparation ===<br />
You must first prepare the skull for implantation. We recommend shaving the head area and sterilizing with 3 alternating betadine and ethanol scrubs. Next, remove the scalp with scissors and clean the skull with hydrogen peroxide and saline. Scrape and score the skull to increase the bond with the glue. We also detach the neck muscle from the skull in order to reduce pull on the skull and prevent muscle growth that can make the implant less stable. Finally we insert a skull screw on the opposite side of the skull to enhance the stability of the implant.<br />
<br />
=== Craniotomy and Aspiration ===<br />
Using a stereotax, align the skull and find the location for the implant. Using a drill, make 4 guide holes to outline the lens placement. For CA1, we recommend offsetting the lens 0.5mm to the medial side of the virus injection to prevent imaging any damage induced by the virus injection. Connect the guide holes to create a circular craniotomy and peel off the circular skull fragment. Do not worry about blood as the top layers of cortex will be removed by aspiration. Make absolutely sure that the craniotomy is at least the size of your GRIN lens! Cover with cortex buffer and scrape around the outside of the craniotomy to remove any excess bone or dura. <br />
<br />
While keeping a constant flow of cortex buffer use the aspirator to slowly remove the cortex above the hippocampus. Continue removing cortex until you reach the white striations of the corpus callosum. Very slowly and carefully remove the horizontal white striations of the corpus callosum until you reach the striations that go vertical. When you reach these striations, stop aspirating and clean up the sides so that the lens can be placed into the brain. Continue washing with cortex buffer until all bleeding stops and keep the brain constantly flushed with cortex buffer.<br />
<br />
=== Lens implantation ===<br />
Attach GRIN lens to lens holder on stereotax. Align the lens with the top of the skull at the most posterior part of the craniotomy and quickly insert the lens 1.35 mm below the top of the skull. Remove any excess liquid with the aspirator. Connect the outside of the GRIN lens with the skull and skull screw using cyanoacrylate glue and let fully dry or use a glue accelerator. Remove the lens holder by simply removing the vacuum suction (you can just pinch the tubing) and withdrawing the holder. Next, cover the entire skull with cyanoacrylate glue, and then cover with dental cement. Let dry completely and cover with Kwik-Sil to protect the lens. Remove from stereotax and allow the animal to recover. Give the animal amoxicillin (or equivalent) for 7 days, and daily injections of carprofen and dexamethasone for 7 days.<br />
<br />
=== Important tips to remember ===<br />
*Try to keep the surgery as sterile as possible. Infections will destroy imaging and the ability to keep the same cells over many days.<br />
<br />
*Try to keep isoflurane as low as possible. Surgeries can take 2-6 hours depending on experience.<br />
<br />
*Aspirations will take a lot of practice. Go slow at first.<br />
<br />
*Depth will be trial and error. If the first few animals don't give good images, check the depth using histology and adjust accordingly.<br />
<br />
<br />
Please feel free to add any tips that you have!<br />
<br />
== Baseplating Protocol ==<br />
<br />
<br />
=== Basic Equipment needed ===<br />
[[File:bp1.png|right|200px|thumb|Put a stack of magnets on a reference Miniscope and mark the top magnet to keep track of its polarity]]<br />
[[File:bp2.png|right|200px|thumb|Magnetically attach the top magnet to screwdriver, coat with cyanoacrylate glue on the edge, and push through hole on baseplate]]<br />
[[File:bp3.png|right|200px|thumb|Repeat for remaining two magnets; ensure top of baseplate is completely flat]]<br />
[[File:bp4.png|right|200px|thumb|Coat bottom of baseplate with cyanoacrylate glue]]<br />
[[File:bp5.png|right|200px|thumb|Score bottom and edges of baseplate with a dental drill]]<br />
You will only need basic surgical equipment to baseplate. This includes a mouse stereotax, an isoflurane vaporizer, surgical heat pad, stereo surgical microscope, light source, and a dental drill. <br />
<br />
You will also need cyanoacrylate glue, fine forceps, double distilled water, lens paper, and dental cement<br />
<br />
=== Baseplate Preparation ===<br />
Before affixing the baseplate on an animal, you will need to prepare the baseplate. <br />
The first step is to insert magnets into the appropriate holes on the baseplate. To ensure that the polarity of the magnets is correct throughout the baseplate preparation, use a reference Miniscope to hold onto the magnets. Mark the top magnet with a sharpie, as seen in the picture to the right. Next, put the top magnet on the head of a screwdriver with the marked side facing away from the screwdriver. Apply cyanoacrylate glue on the edge of the magnet, then push the magnet through the appropriate hole. Ensure that the magnet goes through the hole completely and the top side of the baseplate is flat, as the quality of imaging relies on the Miniscope sitting flush on the baseplate. Repeat this procedure for the next 2 magnets. Finally, put a coat of cyanoacrylate glue on the bottom side of the baseplate and leave the baseplate to dry.<br />
Once the baseplate has dried, score the bottom and the sides of the baseplate using a dental drill to allow the dental glue to adhere the baseplate and skull well.<br />
<br />
=== Cap Preparation ===<br />
Similar to the baseplate preparation, use a reference Miniscope to hold onto the magnets. Mark the top magnet with a sharpie, and put the magnet on the head of a screwdriver with the marked side facing toward the screwdriver. Coat the magnet with cyanoacrylate glue and push the magnet into the appropriate hole. Repeat for the remaining two magnets.<br />
=== Baseplating ===<br />
Connect a Miniscope to a DAQ and initialize the Miniscope software on a computer. Attach your prepared baseplate onto the bottom of the Miniscope using a set screw.<br />
<br />
Using a stereotax and isoflurane vaporizer, anesthetize the animal and fix ear bars until the animal is stable. Carefully remove the Kwik-Sil off the top of the animal’s GRIN lens with blunt forceps being sure not to scratch the lens. Under a stereo surgical microscope, check to see if there is debris on the top of the GRIN lens. If there is, dampen a sheet of lens paper using double distilled water. Wipe the top of the GRIN lens using fine forceps and lens paper, being sure to only wipe laterally and not to apply direct downward pressure.<br />
<br />
After the top of the GRIN lens is clean, explore the field of view with the Miniscope. Adjust the LED brightness and focus accordingly. We advise that you use maximum gain and low LED excitation as to not photobleach the brain during baseplating. After finding your ideal view, practice taking the Miniscope off and on and finding the same view. <br />
<br />
When you feel comfortable repeatedly finding the view with the Miniscope, apply dental cement in a “C” pattern around the GRIN lens, making sure not to touch the lens itself. It will be easier if your first application of dental cement is more viscous. Put the Miniscope on top of the lens and find your ideal view. Hold the Miniscope steadily as the first application of dental cement dries, making sure that your view does not change. <br />
After the first application of dental cement dries, mix more dental cement and apply it on the posterior edge of the baseplate, making sure it seals to the skull. At this point, you can also build up dental cement along the wall of the baseplate on the animal’s right side. Be sure to avoid applying dental cement onto the camera itself and adhering it to the baseplate. Once the dental cement dries and you feel confident that the baseplate will no longer shift, carefully remove the Miniscope from the baseplate by unscrewing the set screw and lifting it off. <br />
<br />
Apply dental cement to the anterior edge of the baseplate and fill in the gaps between the baseplate and skull on the animal’s left side. Allow the dental cement to dry and attach the plastic cap onto the top of the baseplate with a set screw.</div>Chrisl1026http://miniscope.org/index.php?title=Surgery_Protocol&diff=1526Surgery Protocol2016-09-21T04:12:53Z<p>Chrisl1026: /* Basic Equipment needed */</p>
<hr />
<div>The following is a basic outline of our procedure for implanting a GRIN lens above the hippocampus for CA1 imaging. For videos of surgery and baseplating, see the [[Online Workshop]]. Also, see the recent Nature Protocols paper from the Stuber Lab ([http://www.nature.com/nprot/journal/v11/n3/full/nprot.2016.021.html link]) that has a very detailed description of the surgery. <br />
<br />
In order to image, animals will need to undergo three surgical procedures: a virus injection, a GRIN lens implantation, and baseplate attachment.<br />
<br />
== Virus Injection ==<br />
Before implanting the GRIN lens, you will need to inject a fluorescent indicator such as GCaMP6. We have generally used AAV1.Syn.GCaMP6f.WPRE.SV40 from Penn Vector (Item number AV-1-PV2822) and had great success in dorsal CA1. You may also be able to use GCaMP6 transgenic mice, but we have not tested whether any of the newer lines are bright enough to be imaged.<br />
<br />
== GRIN Lens Implantation==<br />
<br />
=== Basic Equipment Needed ===<br />
You will only need basic surgical equipment to perform GRIN lens implantation surgeries. This includes a mouse stereotax, an isoflurane vaporizor, surgical heat pad, stereo surgical microscope, light source, dental drill, and bead sterilizer. For recommended equipment, see the master [https://docs.google.com/spreadsheets/d/12H71DU2QX8d7efUE4yNuikBEiIzKaXjYqdc0A-oLNSw/edit?usp=sharing parts list].<br />
<br />
You will need the following basic tools and supplies: fine forceps, blunt forceps, fine scissors, scalpel and blade, small skull screws, drill bits, cyanoacrylate glue, dental cement, Kwik-Sil, and [[cortex buffer]]<br />
<br />
=== Aspirator and Lens Holder ===<br />
You will need an aspirator to remove cortex above the hippocampus. This can be built very simply with a vacuum line (or pump), a liquid trap, 1 ml syringe (with a hole to control suction), blunt needles, tubing and connectors, and hot glue.<br />
[[File:Aspirator.PNG|center|500px]]<br />
<br />
You can easily build a GRIN lens holder by using a suction system to hold the lens in place as it is dropped into the brain for implantation. To build, start with two 1ml micropipette tips. Cut the first so that the tip is just larger than the diameter of the GRIN lens you will be using. Cut the second tip so that the end is just smaller than the size of the GRIN lens. Connect this end to a 1ml syringe and connect to a vacuum line. This will then hold the lens in place during implantation. You can then use tape to attach this holder to a stereotax. Another solution we have used is a basic drill chuck, but this is only good for large (2mm) lenses.<br />
[[File:LensHolder.PNG|center|500px]]<br />
<br />
=== Skull Preparation ===<br />
You must first prepare the skull for implantation. We recommend shaving the head area and sterilizing with 3 alternating betadine and ethanol scrubs. Next, remove the scalp with scissors and clean the skull with hydrogen peroxide and saline. Scrape and score the skull to increase the bond with the glue. We also detach the neck muscle from the skull in order to reduce pull on the skull and prevent muscle growth that can make the implant less stable. Finally we insert a skull screw on the opposite side of the skull to enhance the stability of the implant.<br />
<br />
=== Craniotomy and Aspiration ===<br />
Using a stereotax, align the skull and find the location for the implant. Using a drill, make 4 guide holes to outline the lens placement. For CA1, we recommend offsetting the lens 0.5mm to the medial side of the virus injection to prevent imaging any damage induced by the virus injection. Connect the guide holes to create a circular craniotomy and peel off the circular skull fragment. Do not worry about blood as the top layers of cortex will be removed by aspiration. Make absolutely sure that the craniotomy is at least the size of your GRIN lens! Cover with cortex buffer and scrape around the outside of the craniotomy to remove any excess bone or dura. <br />
<br />
While keeping a constant flow of cortex buffer use the aspirator to slowly remove the cortex above the hippocampus. Continue removing cortex until you reach the white striations of the corpus callosum. Very slowly and carefully remove the horizontal white striations of the corpus callosum until you reach the striations that go vertical. When you reach these striations, stop aspirating and clean up the sides so that the lens can be placed into the brain. Continue washing with cortex buffer until all bleeding stops and keep the brain constantly flushed with cortex buffer.<br />
<br />
=== Lens implantation ===<br />
Attach GRIN lens to lens holder on stereotax. Align the lens with the top of the skull at the most posterior part of the craniotomy and quickly insert the lens 1.35 mm below the top of the skull. Remove any excess liquid with the aspirator. Connect the outside of the GRIN lens with the skull and skull screw using cyanoacrylate glue and let fully dry or use a glue accelerator. Remove the lens holder by simply removing the vacuum suction (you can just pinch the tubing) and withdrawing the holder. Next, cover the entire skull with cyanoacrylate glue, and then cover with dental cement. Let dry completely and cover with Kwik-Sil to protect the lens. Remove from stereotax and allow the animal to recover. Give the animal amoxicillin (or equivalent) for 7 days, and daily injections of carprofen and dexamethasone for 7 days.<br />
<br />
=== Important tips to remember ===<br />
*Try to keep the surgery as sterile as possible. Infections will destroy imaging and the ability to keep the same cells over many days.<br />
<br />
*Try to keep isoflurane as low as possible. Surgeries can take 2-6 hours depending on experience.<br />
<br />
*Aspirations will take a lot of practice. Go slow at first.<br />
<br />
*Depth will be trial and error. If the first few animals don't give good images, check the depth using histology and adjust accordingly.<br />
<br />
<br />
Please feel free to add any tips that you have!<br />
<br />
== Baseplating Protocol ==<br />
<br />
<br />
=== Basic Equipment needed ===<br />
You will only need basic surgical equipment to baseplate. This includes a mouse stereotax, an isoflurane vaporizer, surgical heat pad, stereo surgical microscope, light source, and a dental drill. <br />
<br />
You will also need cyanoacrylate glue, fine forceps, double distilled water, lens paper, and dental cement<br />
[[File:bp1.png|right|200px|thumb|Put a stack of magnets on a reference Miniscope and mark the top magnet to keep track of its polarity]]<br />
[[File:bp2.png|right|200px|thumb|Magnetically attach the top magnet to screwdriver, coat with cyanoacrylate glue on the edge, and push through hole on baseplate]]<br />
[[File:bp3.png|right|200px|thumb|Repeat for remaining two magnets; ensure top of baseplate is completely flat]]<br />
[[File:bp4.png|right|200px|thumb|Coat bottom of baseplate with cyanoacrylate glue]]<br />
[[File:bp5.png|right|200px|thumb|Score bottom and edges of baseplate with a dental drill]]<br />
<br />
=== Baseplate Preparation ===<br />
Before affixing the baseplate on an animal, you will need to prepare the baseplate. <br />
The first step is to insert magnets into the appropriate holes on the baseplate. To ensure that the polarity of the magnets is correct throughout the baseplate preparation, use a reference Miniscope to hold onto the magnets. Mark the top magnet with a sharpie, as seen in the picture to the right. Next, put the top magnet on the head of a screwdriver with the marked side facing away from the screwdriver. Apply cyanoacrylate glue on the edge of the magnet, then push the magnet through the appropriate hole. Ensure that the magnet goes through the hole completely and the top side of the baseplate is flat, as the quality of imaging relies on the Miniscope sitting flush on the baseplate. Repeat this procedure for the next 2 magnets. Finally, put a coat of cyanoacrylate glue on the bottom side of the baseplate and leave the baseplate to dry.<br />
Once the baseplate has dried, score the bottom and the sides of the baseplate using a dental drill to allow the dental glue to adhere the baseplate and skull well.<br />
<br />
=== Cap Preparation ===<br />
Similar to the baseplate preparation, use a reference Miniscope to hold onto the magnets. Mark the top magnet with a sharpie, and put the magnet on the head of a screwdriver with the marked side facing toward the screwdriver. Coat the magnet with cyanoacrylate glue and push the magnet into the appropriate hole. Repeat for the remaining two magnets.<br />
=== Baseplating ===<br />
Connect a Miniscope to a DAQ and initialize the Miniscope software on a computer. Attach your prepared baseplate onto the bottom of the Miniscope using a set screw.<br />
<br />
Using a stereotax and isoflurane vaporizer, anesthetize the animal and fix ear bars until the animal is stable. Carefully remove the Kwik-Sil off the top of the animal’s GRIN lens with blunt forceps being sure not to scratch the lens. Under a stereo surgical microscope, check to see if there is debris on the top of the GRIN lens. If there is, dampen a sheet of lens paper using double distilled water. Wipe the top of the GRIN lens using fine forceps and lens paper, being sure to only wipe laterally and not to apply direct downward pressure.<br />
<br />
After the top of the GRIN lens is clean, explore the field of view with the Miniscope. Adjust the LED brightness and focus accordingly. We advise that you use maximum gain and low LED excitation as to not photobleach the brain during baseplating. After finding your ideal view, practice taking the Miniscope off and on and finding the same view. <br />
<br />
When you feel comfortable repeatedly finding the view with the Miniscope, apply dental cement in a “C” pattern around the GRIN lens, making sure not to touch the lens itself. It will be easier if your first application of dental cement is more viscous. Put the Miniscope on top of the lens and find your ideal view. Hold the Miniscope steadily as the first application of dental cement dries, making sure that your view does not change. <br />
After the first application of dental cement dries, mix more dental cement and apply it on the posterior edge of the baseplate, making sure it seals to the skull. At this point, you can also build up dental cement along the wall of the baseplate on the animal’s right side. Be sure to avoid applying dental cement onto the camera itself and adhering it to the baseplate. Once the dental cement dries and you feel confident that the baseplate will no longer shift, carefully remove the Miniscope from the baseplate by unscrewing the set screw and lifting it off. <br />
<br />
Apply dental cement to the anterior edge of the baseplate and fill in the gaps between the baseplate and skull on the animal’s left side. Allow the dental cement to dry and attach the plastic cap onto the top of the baseplate with a set screw.</div>Chrisl1026http://miniscope.org/index.php?title=Surgery_Protocol&diff=1525Surgery Protocol2016-09-21T04:12:39Z<p>Chrisl1026: /* Baseplate Preparation */</p>
<hr />
<div>The following is a basic outline of our procedure for implanting a GRIN lens above the hippocampus for CA1 imaging. For videos of surgery and baseplating, see the [[Online Workshop]]. Also, see the recent Nature Protocols paper from the Stuber Lab ([http://www.nature.com/nprot/journal/v11/n3/full/nprot.2016.021.html link]) that has a very detailed description of the surgery. <br />
<br />
In order to image, animals will need to undergo three surgical procedures: a virus injection, a GRIN lens implantation, and baseplate attachment.<br />
<br />
== Virus Injection ==<br />
Before implanting the GRIN lens, you will need to inject a fluorescent indicator such as GCaMP6. We have generally used AAV1.Syn.GCaMP6f.WPRE.SV40 from Penn Vector (Item number AV-1-PV2822) and had great success in dorsal CA1. You may also be able to use GCaMP6 transgenic mice, but we have not tested whether any of the newer lines are bright enough to be imaged.<br />
<br />
== GRIN Lens Implantation==<br />
<br />
=== Basic Equipment Needed ===<br />
You will only need basic surgical equipment to perform GRIN lens implantation surgeries. This includes a mouse stereotax, an isoflurane vaporizor, surgical heat pad, stereo surgical microscope, light source, dental drill, and bead sterilizer. For recommended equipment, see the master [https://docs.google.com/spreadsheets/d/12H71DU2QX8d7efUE4yNuikBEiIzKaXjYqdc0A-oLNSw/edit?usp=sharing parts list].<br />
<br />
You will need the following basic tools and supplies: fine forceps, blunt forceps, fine scissors, scalpel and blade, small skull screws, drill bits, cyanoacrylate glue, dental cement, Kwik-Sil, and [[cortex buffer]]<br />
<br />
=== Aspirator and Lens Holder ===<br />
You will need an aspirator to remove cortex above the hippocampus. This can be built very simply with a vacuum line (or pump), a liquid trap, 1 ml syringe (with a hole to control suction), blunt needles, tubing and connectors, and hot glue.<br />
[[File:Aspirator.PNG|center|500px]]<br />
<br />
You can easily build a GRIN lens holder by using a suction system to hold the lens in place as it is dropped into the brain for implantation. To build, start with two 1ml micropipette tips. Cut the first so that the tip is just larger than the diameter of the GRIN lens you will be using. Cut the second tip so that the end is just smaller than the size of the GRIN lens. Connect this end to a 1ml syringe and connect to a vacuum line. This will then hold the lens in place during implantation. You can then use tape to attach this holder to a stereotax. Another solution we have used is a basic drill chuck, but this is only good for large (2mm) lenses.<br />
[[File:LensHolder.PNG|center|500px]]<br />
<br />
=== Skull Preparation ===<br />
You must first prepare the skull for implantation. We recommend shaving the head area and sterilizing with 3 alternating betadine and ethanol scrubs. Next, remove the scalp with scissors and clean the skull with hydrogen peroxide and saline. Scrape and score the skull to increase the bond with the glue. We also detach the neck muscle from the skull in order to reduce pull on the skull and prevent muscle growth that can make the implant less stable. Finally we insert a skull screw on the opposite side of the skull to enhance the stability of the implant.<br />
<br />
=== Craniotomy and Aspiration ===<br />
Using a stereotax, align the skull and find the location for the implant. Using a drill, make 4 guide holes to outline the lens placement. For CA1, we recommend offsetting the lens 0.5mm to the medial side of the virus injection to prevent imaging any damage induced by the virus injection. Connect the guide holes to create a circular craniotomy and peel off the circular skull fragment. Do not worry about blood as the top layers of cortex will be removed by aspiration. Make absolutely sure that the craniotomy is at least the size of your GRIN lens! Cover with cortex buffer and scrape around the outside of the craniotomy to remove any excess bone or dura. <br />
<br />
While keeping a constant flow of cortex buffer use the aspirator to slowly remove the cortex above the hippocampus. Continue removing cortex until you reach the white striations of the corpus callosum. Very slowly and carefully remove the horizontal white striations of the corpus callosum until you reach the striations that go vertical. When you reach these striations, stop aspirating and clean up the sides so that the lens can be placed into the brain. Continue washing with cortex buffer until all bleeding stops and keep the brain constantly flushed with cortex buffer.<br />
<br />
=== Lens implantation ===<br />
Attach GRIN lens to lens holder on stereotax. Align the lens with the top of the skull at the most posterior part of the craniotomy and quickly insert the lens 1.35 mm below the top of the skull. Remove any excess liquid with the aspirator. Connect the outside of the GRIN lens with the skull and skull screw using cyanoacrylate glue and let fully dry or use a glue accelerator. Remove the lens holder by simply removing the vacuum suction (you can just pinch the tubing) and withdrawing the holder. Next, cover the entire skull with cyanoacrylate glue, and then cover with dental cement. Let dry completely and cover with Kwik-Sil to protect the lens. Remove from stereotax and allow the animal to recover. Give the animal amoxicillin (or equivalent) for 7 days, and daily injections of carprofen and dexamethasone for 7 days.<br />
<br />
=== Important tips to remember ===<br />
*Try to keep the surgery as sterile as possible. Infections will destroy imaging and the ability to keep the same cells over many days.<br />
<br />
*Try to keep isoflurane as low as possible. Surgeries can take 2-6 hours depending on experience.<br />
<br />
*Aspirations will take a lot of practice. Go slow at first.<br />
<br />
*Depth will be trial and error. If the first few animals don't give good images, check the depth using histology and adjust accordingly.<br />
<br />
<br />
Please feel free to add any tips that you have!<br />
<br />
== Baseplating Protocol ==<br />
<br />
<br />
=== Basic Equipment needed ===<br />
You will only need basic surgical equipment to baseplate. This includes a mouse stereotax, an isoflurane vaporizer, surgical heat pad, stereo surgical microscope, light source, and a dental drill. <br />
<br />
You will also need cyanoacrylate glue, fine forceps, double distilled water, lens paper, and dental cement<br />
=== Baseplate Preparation ===<br />
Before affixing the baseplate on an animal, you will need to prepare the baseplate. <br />
The first step is to insert magnets into the appropriate holes on the baseplate. To ensure that the polarity of the magnets is correct throughout the baseplate preparation, use a reference Miniscope to hold onto the magnets. Mark the top magnet with a sharpie, as seen in the picture to the right. Next, put the top magnet on the head of a screwdriver with the marked side facing away from the screwdriver. Apply cyanoacrylate glue on the edge of the magnet, then push the magnet through the appropriate hole. Ensure that the magnet goes through the hole completely and the top side of the baseplate is flat, as the quality of imaging relies on the Miniscope sitting flush on the baseplate. Repeat this procedure for the next 2 magnets. Finally, put a coat of cyanoacrylate glue on the bottom side of the baseplate and leave the baseplate to dry.<br />
Once the baseplate has dried, score the bottom and the sides of the baseplate using a dental drill to allow the dental glue to adhere the baseplate and skull well.<br />
<br />
=== Cap Preparation ===<br />
Similar to the baseplate preparation, use a reference Miniscope to hold onto the magnets. Mark the top magnet with a sharpie, and put the magnet on the head of a screwdriver with the marked side facing toward the screwdriver. Coat the magnet with cyanoacrylate glue and push the magnet into the appropriate hole. Repeat for the remaining two magnets.<br />
=== Baseplating ===<br />
Connect a Miniscope to a DAQ and initialize the Miniscope software on a computer. Attach your prepared baseplate onto the bottom of the Miniscope using a set screw.<br />
<br />
Using a stereotax and isoflurane vaporizer, anesthetize the animal and fix ear bars until the animal is stable. Carefully remove the Kwik-Sil off the top of the animal’s GRIN lens with blunt forceps being sure not to scratch the lens. Under a stereo surgical microscope, check to see if there is debris on the top of the GRIN lens. If there is, dampen a sheet of lens paper using double distilled water. Wipe the top of the GRIN lens using fine forceps and lens paper, being sure to only wipe laterally and not to apply direct downward pressure.<br />
<br />
After the top of the GRIN lens is clean, explore the field of view with the Miniscope. Adjust the LED brightness and focus accordingly. We advise that you use maximum gain and low LED excitation as to not photobleach the brain during baseplating. After finding your ideal view, practice taking the Miniscope off and on and finding the same view. <br />
<br />
When you feel comfortable repeatedly finding the view with the Miniscope, apply dental cement in a “C” pattern around the GRIN lens, making sure not to touch the lens itself. It will be easier if your first application of dental cement is more viscous. Put the Miniscope on top of the lens and find your ideal view. Hold the Miniscope steadily as the first application of dental cement dries, making sure that your view does not change. <br />
After the first application of dental cement dries, mix more dental cement and apply it on the posterior edge of the baseplate, making sure it seals to the skull. At this point, you can also build up dental cement along the wall of the baseplate on the animal’s right side. Be sure to avoid applying dental cement onto the camera itself and adhering it to the baseplate. Once the dental cement dries and you feel confident that the baseplate will no longer shift, carefully remove the Miniscope from the baseplate by unscrewing the set screw and lifting it off. <br />
<br />
Apply dental cement to the anterior edge of the baseplate and fill in the gaps between the baseplate and skull on the animal’s left side. Allow the dental cement to dry and attach the plastic cap onto the top of the baseplate with a set screw.</div>Chrisl1026http://miniscope.org/index.php?title=Surgery_Protocol&diff=1524Surgery Protocol2016-09-21T04:00:05Z<p>Chrisl1026: /* Baseplate Preparation */</p>
<hr />
<div>The following is a basic outline of our procedure for implanting a GRIN lens above the hippocampus for CA1 imaging. For videos of surgery and baseplating, see the [[Online Workshop]]. Also, see the recent Nature Protocols paper from the Stuber Lab ([http://www.nature.com/nprot/journal/v11/n3/full/nprot.2016.021.html link]) that has a very detailed description of the surgery. <br />
<br />
In order to image, animals will need to undergo three surgical procedures: a virus injection, a GRIN lens implantation, and baseplate attachment.<br />
<br />
== Virus Injection ==<br />
Before implanting the GRIN lens, you will need to inject a fluorescent indicator such as GCaMP6. We have generally used AAV1.Syn.GCaMP6f.WPRE.SV40 from Penn Vector (Item number AV-1-PV2822) and had great success in dorsal CA1. You may also be able to use GCaMP6 transgenic mice, but we have not tested whether any of the newer lines are bright enough to be imaged.<br />
<br />
== GRIN Lens Implantation==<br />
<br />
=== Basic Equipment Needed ===<br />
You will only need basic surgical equipment to perform GRIN lens implantation surgeries. This includes a mouse stereotax, an isoflurane vaporizor, surgical heat pad, stereo surgical microscope, light source, dental drill, and bead sterilizer. For recommended equipment, see the master [https://docs.google.com/spreadsheets/d/12H71DU2QX8d7efUE4yNuikBEiIzKaXjYqdc0A-oLNSw/edit?usp=sharing parts list].<br />
<br />
You will need the following basic tools and supplies: fine forceps, blunt forceps, fine scissors, scalpel and blade, small skull screws, drill bits, cyanoacrylate glue, dental cement, Kwik-Sil, and [[cortex buffer]]<br />
<br />
=== Aspirator and Lens Holder ===<br />
You will need an aspirator to remove cortex above the hippocampus. This can be built very simply with a vacuum line (or pump), a liquid trap, 1 ml syringe (with a hole to control suction), blunt needles, tubing and connectors, and hot glue.<br />
[[File:Aspirator.PNG|center|500px]]<br />
<br />
You can easily build a GRIN lens holder by using a suction system to hold the lens in place as it is dropped into the brain for implantation. To build, start with two 1ml micropipette tips. Cut the first so that the tip is just larger than the diameter of the GRIN lens you will be using. Cut the second tip so that the end is just smaller than the size of the GRIN lens. Connect this end to a 1ml syringe and connect to a vacuum line. This will then hold the lens in place during implantation. You can then use tape to attach this holder to a stereotax. Another solution we have used is a basic drill chuck, but this is only good for large (2mm) lenses.<br />
[[File:LensHolder.PNG|center|500px]]<br />
<br />
=== Skull Preparation ===<br />
You must first prepare the skull for implantation. We recommend shaving the head area and sterilizing with 3 alternating betadine and ethanol scrubs. Next, remove the scalp with scissors and clean the skull with hydrogen peroxide and saline. Scrape and score the skull to increase the bond with the glue. We also detach the neck muscle from the skull in order to reduce pull on the skull and prevent muscle growth that can make the implant less stable. Finally we insert a skull screw on the opposite side of the skull to enhance the stability of the implant.<br />
<br />
=== Craniotomy and Aspiration ===<br />
Using a stereotax, align the skull and find the location for the implant. Using a drill, make 4 guide holes to outline the lens placement. For CA1, we recommend offsetting the lens 0.5mm to the medial side of the virus injection to prevent imaging any damage induced by the virus injection. Connect the guide holes to create a circular craniotomy and peel off the circular skull fragment. Do not worry about blood as the top layers of cortex will be removed by aspiration. Make absolutely sure that the craniotomy is at least the size of your GRIN lens! Cover with cortex buffer and scrape around the outside of the craniotomy to remove any excess bone or dura. <br />
<br />
While keeping a constant flow of cortex buffer use the aspirator to slowly remove the cortex above the hippocampus. Continue removing cortex until you reach the white striations of the corpus callosum. Very slowly and carefully remove the horizontal white striations of the corpus callosum until you reach the striations that go vertical. When you reach these striations, stop aspirating and clean up the sides so that the lens can be placed into the brain. Continue washing with cortex buffer until all bleeding stops and keep the brain constantly flushed with cortex buffer.<br />
<br />
=== Lens implantation ===<br />
Attach GRIN lens to lens holder on stereotax. Align the lens with the top of the skull at the most posterior part of the craniotomy and quickly insert the lens 1.35 mm below the top of the skull. Remove any excess liquid with the aspirator. Connect the outside of the GRIN lens with the skull and skull screw using cyanoacrylate glue and let fully dry or use a glue accelerator. Remove the lens holder by simply removing the vacuum suction (you can just pinch the tubing) and withdrawing the holder. Next, cover the entire skull with cyanoacrylate glue, and then cover with dental cement. Let dry completely and cover with Kwik-Sil to protect the lens. Remove from stereotax and allow the animal to recover. Give the animal amoxicillin (or equivalent) for 7 days, and daily injections of carprofen and dexamethasone for 7 days.<br />
<br />
=== Important tips to remember ===<br />
*Try to keep the surgery as sterile as possible. Infections will destroy imaging and the ability to keep the same cells over many days.<br />
<br />
*Try to keep isoflurane as low as possible. Surgeries can take 2-6 hours depending on experience.<br />
<br />
*Aspirations will take a lot of practice. Go slow at first.<br />
<br />
*Depth will be trial and error. If the first few animals don't give good images, check the depth using histology and adjust accordingly.<br />
<br />
<br />
Please feel free to add any tips that you have!<br />
<br />
== Baseplating Protocol ==<br />
<br />
<br />
=== Basic Equipment needed ===<br />
You will only need basic surgical equipment to baseplate. This includes a mouse stereotax, an isoflurane vaporizer, surgical heat pad, stereo surgical microscope, light source, and a dental drill. <br />
<br />
You will also need cyanoacrylate glue, fine forceps, double distilled water, lens paper, and dental cement<br />
=== Baseplate Preparation ===<br />
Before affixing the baseplate on an animal, you will need to prepare the baseplate. [[File:bp1.png|right|200px]]<br />
[[File:bp2.png|right|200px]]<br />
The first step is to insert magnets into the appropriate holes on the baseplate. To ensure that the polarity of the magnets is correct throughout the baseplate preparation, use a reference Miniscope to hold onto the magnets. Mark the top magnet with a sharpie, as seen in the picture to the right. Next, put the top magnet on the head of a screwdriver with the marked side facing away from the screwdriver. Apply cyanoacrylate glue on the edge of the magnet, then push the magnet through the appropriate hole. Ensure that the magnet goes through the hole completely and the top side of the baseplate is flat, as the quality of imaging relies on the Miniscope sitting flush on the baseplate. Repeat this procedure for the next 2 magnets. Finally, put a coat of cyanoacrylate glue on the bottom side of the baseplate and leave the baseplate to dry.[[File:bp3.png|right|200px]]<br />
[[File:bp4.png|right|200px]]<br />
Once the baseplate has dried, score the bottom and the sides of the baseplate using a dental drill to allow the dental glue to adhere the baseplate and skull well.<br />
[[File:bp5.png|right|200px]]<br />
<br />
=== Cap Preparation ===<br />
Similar to the baseplate preparation, use a reference Miniscope to hold onto the magnets. Mark the top magnet with a sharpie, and put the magnet on the head of a screwdriver with the marked side facing toward the screwdriver. Coat the magnet with cyanoacrylate glue and push the magnet into the appropriate hole. Repeat for the remaining two magnets.<br />
=== Baseplating ===<br />
Connect a Miniscope to a DAQ and initialize the Miniscope software on a computer. Attach your prepared baseplate onto the bottom of the Miniscope using a set screw.<br />
<br />
Using a stereotax and isoflurane vaporizer, anesthetize the animal and fix ear bars until the animal is stable. Carefully remove the Kwik-Sil off the top of the animal’s GRIN lens with blunt forceps being sure not to scratch the lens. Under a stereo surgical microscope, check to see if there is debris on the top of the GRIN lens. If there is, dampen a sheet of lens paper using double distilled water. Wipe the top of the GRIN lens using fine forceps and lens paper, being sure to only wipe laterally and not to apply direct downward pressure.<br />
<br />
After the top of the GRIN lens is clean, explore the field of view with the Miniscope. Adjust the LED brightness and focus accordingly. We advise that you use maximum gain and low LED excitation as to not photobleach the brain during baseplating. After finding your ideal view, practice taking the Miniscope off and on and finding the same view. <br />
<br />
When you feel comfortable repeatedly finding the view with the Miniscope, apply dental cement in a “C” pattern around the GRIN lens, making sure not to touch the lens itself. It will be easier if your first application of dental cement is more viscous. Put the Miniscope on top of the lens and find your ideal view. Hold the Miniscope steadily as the first application of dental cement dries, making sure that your view does not change. <br />
After the first application of dental cement dries, mix more dental cement and apply it on the posterior edge of the baseplate, making sure it seals to the skull. At this point, you can also build up dental cement along the wall of the baseplate on the animal’s right side. Be sure to avoid applying dental cement onto the camera itself and adhering it to the baseplate. Once the dental cement dries and you feel confident that the baseplate will no longer shift, carefully remove the Miniscope from the baseplate by unscrewing the set screw and lifting it off. <br />
<br />
Apply dental cement to the anterior edge of the baseplate and fill in the gaps between the baseplate and skull on the animal’s left side. Allow the dental cement to dry and attach the plastic cap onto the top of the baseplate with a set screw.</div>Chrisl1026http://miniscope.org/index.php?title=Surgery_Protocol&diff=1523Surgery Protocol2016-09-21T03:54:02Z<p>Chrisl1026: /* Baseplate Preparation */</p>
<hr />
<div>The following is a basic outline of our procedure for implanting a GRIN lens above the hippocampus for CA1 imaging. For videos of surgery and baseplating, see the [[Online Workshop]]. Also, see the recent Nature Protocols paper from the Stuber Lab ([http://www.nature.com/nprot/journal/v11/n3/full/nprot.2016.021.html link]) that has a very detailed description of the surgery. <br />
<br />
In order to image, animals will need to undergo three surgical procedures: a virus injection, a GRIN lens implantation, and baseplate attachment.<br />
<br />
== Virus Injection ==<br />
Before implanting the GRIN lens, you will need to inject a fluorescent indicator such as GCaMP6. We have generally used AAV1.Syn.GCaMP6f.WPRE.SV40 from Penn Vector (Item number AV-1-PV2822) and had great success in dorsal CA1. You may also be able to use GCaMP6 transgenic mice, but we have not tested whether any of the newer lines are bright enough to be imaged.<br />
<br />
== GRIN Lens Implantation==<br />
<br />
=== Basic Equipment Needed ===<br />
You will only need basic surgical equipment to perform GRIN lens implantation surgeries. This includes a mouse stereotax, an isoflurane vaporizor, surgical heat pad, stereo surgical microscope, light source, dental drill, and bead sterilizer. For recommended equipment, see the master [https://docs.google.com/spreadsheets/d/12H71DU2QX8d7efUE4yNuikBEiIzKaXjYqdc0A-oLNSw/edit?usp=sharing parts list].<br />
<br />
You will need the following basic tools and supplies: fine forceps, blunt forceps, fine scissors, scalpel and blade, small skull screws, drill bits, cyanoacrylate glue, dental cement, Kwik-Sil, and [[cortex buffer]]<br />
<br />
=== Aspirator and Lens Holder ===<br />
You will need an aspirator to remove cortex above the hippocampus. This can be built very simply with a vacuum line (or pump), a liquid trap, 1 ml syringe (with a hole to control suction), blunt needles, tubing and connectors, and hot glue.<br />
[[File:Aspirator.PNG|center|500px]]<br />
<br />
You can easily build a GRIN lens holder by using a suction system to hold the lens in place as it is dropped into the brain for implantation. To build, start with two 1ml micropipette tips. Cut the first so that the tip is just larger than the diameter of the GRIN lens you will be using. Cut the second tip so that the end is just smaller than the size of the GRIN lens. Connect this end to a 1ml syringe and connect to a vacuum line. This will then hold the lens in place during implantation. You can then use tape to attach this holder to a stereotax. Another solution we have used is a basic drill chuck, but this is only good for large (2mm) lenses.<br />
[[File:LensHolder.PNG|center|500px]]<br />
<br />
=== Skull Preparation ===<br />
You must first prepare the skull for implantation. We recommend shaving the head area and sterilizing with 3 alternating betadine and ethanol scrubs. Next, remove the scalp with scissors and clean the skull with hydrogen peroxide and saline. Scrape and score the skull to increase the bond with the glue. We also detach the neck muscle from the skull in order to reduce pull on the skull and prevent muscle growth that can make the implant less stable. Finally we insert a skull screw on the opposite side of the skull to enhance the stability of the implant.<br />
<br />
=== Craniotomy and Aspiration ===<br />
Using a stereotax, align the skull and find the location for the implant. Using a drill, make 4 guide holes to outline the lens placement. For CA1, we recommend offsetting the lens 0.5mm to the medial side of the virus injection to prevent imaging any damage induced by the virus injection. Connect the guide holes to create a circular craniotomy and peel off the circular skull fragment. Do not worry about blood as the top layers of cortex will be removed by aspiration. Make absolutely sure that the craniotomy is at least the size of your GRIN lens! Cover with cortex buffer and scrape around the outside of the craniotomy to remove any excess bone or dura. <br />
<br />
While keeping a constant flow of cortex buffer use the aspirator to slowly remove the cortex above the hippocampus. Continue removing cortex until you reach the white striations of the corpus callosum. Very slowly and carefully remove the horizontal white striations of the corpus callosum until you reach the striations that go vertical. When you reach these striations, stop aspirating and clean up the sides so that the lens can be placed into the brain. Continue washing with cortex buffer until all bleeding stops and keep the brain constantly flushed with cortex buffer.<br />
<br />
=== Lens implantation ===<br />
Attach GRIN lens to lens holder on stereotax. Align the lens with the top of the skull at the most posterior part of the craniotomy and quickly insert the lens 1.35 mm below the top of the skull. Remove any excess liquid with the aspirator. Connect the outside of the GRIN lens with the skull and skull screw using cyanoacrylate glue and let fully dry or use a glue accelerator. Remove the lens holder by simply removing the vacuum suction (you can just pinch the tubing) and withdrawing the holder. Next, cover the entire skull with cyanoacrylate glue, and then cover with dental cement. Let dry completely and cover with Kwik-Sil to protect the lens. Remove from stereotax and allow the animal to recover. Give the animal amoxicillin (or equivalent) for 7 days, and daily injections of carprofen and dexamethasone for 7 days.<br />
<br />
=== Important tips to remember ===<br />
*Try to keep the surgery as sterile as possible. Infections will destroy imaging and the ability to keep the same cells over many days.<br />
<br />
*Try to keep isoflurane as low as possible. Surgeries can take 2-6 hours depending on experience.<br />
<br />
*Aspirations will take a lot of practice. Go slow at first.<br />
<br />
*Depth will be trial and error. If the first few animals don't give good images, check the depth using histology and adjust accordingly.<br />
<br />
<br />
Please feel free to add any tips that you have!<br />
<br />
== Baseplating Protocol ==<br />
<br />
<br />
=== Basic Equipment needed ===<br />
You will only need basic surgical equipment to baseplate. This includes a mouse stereotax, an isoflurane vaporizer, surgical heat pad, stereo surgical microscope, light source, and a dental drill. <br />
<br />
You will also need cyanoacrylate glue, fine forceps, double distilled water, lens paper, and dental cement<br />
=== Baseplate Preparation ===<br />
Before affixing the baseplate on an animal, you will need to prepare the baseplate. [[File:bp1.png|right|200px]]<br />
[[File:bp2.png|right|200px]]<br />
The first step is to insert magnets into the appropriate holes on the baseplate. To ensure that the polarity of the magnets is correct throughout the baseplate preparation, use a reference Miniscope to hold onto the magnets. Mark the top magnet with a sharpie, as seen in the picture below. Next, put the top magnet on the head of a screwdriver with the marked side facing away from the screwdriver. Apply cyanoacrylate glue on the edge of the magnet, then push the magnet through the appropriate hole. Ensure that the magnet goes through the hole completely and the top side of the baseplate is flat, as the quality of imaging relies on the Miniscope sitting flush on the baseplate. Repeat this procedure for the next 2 magnets. Finally, put a coat of cyanoacrylate glue on the bottom side of the baseplate and leave the baseplate to dry.[[File:bp3.png|right|200px]]<br />
[[File:bp4.png|right|200px]]<br />
Once the baseplate has dried, score the bottom and the sides of the baseplate using a dental drill to allow the dental glue to adhere the baseplate and skull well.<br />
[[File:bp5.png|right|200px]]<br />
<br />
=== Cap Preparation ===<br />
Similar to the baseplate preparation, use a reference Miniscope to hold onto the magnets. Mark the top magnet with a sharpie, and put the magnet on the head of a screwdriver with the marked side facing toward the screwdriver. Coat the magnet with cyanoacrylate glue and push the magnet into the appropriate hole. Repeat for the remaining two magnets.<br />
=== Baseplating ===<br />
Connect a Miniscope to a DAQ and initialize the Miniscope software on a computer. Attach your prepared baseplate onto the bottom of the Miniscope using a set screw.<br />
<br />
Using a stereotax and isoflurane vaporizer, anesthetize the animal and fix ear bars until the animal is stable. Carefully remove the Kwik-Sil off the top of the animal’s GRIN lens with blunt forceps being sure not to scratch the lens. Under a stereo surgical microscope, check to see if there is debris on the top of the GRIN lens. If there is, dampen a sheet of lens paper using double distilled water. Wipe the top of the GRIN lens using fine forceps and lens paper, being sure to only wipe laterally and not to apply direct downward pressure.<br />
<br />
After the top of the GRIN lens is clean, explore the field of view with the Miniscope. Adjust the LED brightness and focus accordingly. We advise that you use maximum gain and low LED excitation as to not photobleach the brain during baseplating. After finding your ideal view, practice taking the Miniscope off and on and finding the same view. <br />
<br />
When you feel comfortable repeatedly finding the view with the Miniscope, apply dental cement in a “C” pattern around the GRIN lens, making sure not to touch the lens itself. It will be easier if your first application of dental cement is more viscous. Put the Miniscope on top of the lens and find your ideal view. Hold the Miniscope steadily as the first application of dental cement dries, making sure that your view does not change. <br />
After the first application of dental cement dries, mix more dental cement and apply it on the posterior edge of the baseplate, making sure it seals to the skull. At this point, you can also build up dental cement along the wall of the baseplate on the animal’s right side. Be sure to avoid applying dental cement onto the camera itself and adhering it to the baseplate. Once the dental cement dries and you feel confident that the baseplate will no longer shift, carefully remove the Miniscope from the baseplate by unscrewing the set screw and lifting it off. <br />
<br />
Apply dental cement to the anterior edge of the baseplate and fill in the gaps between the baseplate and skull on the animal’s left side. Allow the dental cement to dry and attach the plastic cap onto the top of the baseplate with a set screw.</div>Chrisl1026http://miniscope.org/index.php?title=File:Bp5.png&diff=1522File:Bp5.png2016-09-21T03:49:24Z<p>Chrisl1026: </p>
<hr />
<div></div>Chrisl1026http://miniscope.org/index.php?title=File:Bp4.png&diff=1521File:Bp4.png2016-09-21T03:48:40Z<p>Chrisl1026: </p>
<hr />
<div></div>Chrisl1026http://miniscope.org/index.php?title=File:Bp3.png&diff=1520File:Bp3.png2016-09-21T03:47:56Z<p>Chrisl1026: </p>
<hr />
<div></div>Chrisl1026http://miniscope.org/index.php?title=File:Bp2.png&diff=1519File:Bp2.png2016-09-21T03:47:24Z<p>Chrisl1026: </p>
<hr />
<div></div>Chrisl1026http://miniscope.org/index.php?title=File:Bp1.png&diff=1518File:Bp1.png2016-09-21T03:46:53Z<p>Chrisl1026: </p>
<hr />
<div></div>Chrisl1026http://miniscope.org/index.php?title=Surgery_Protocol&diff=1517Surgery Protocol2016-09-21T03:42:26Z<p>Chrisl1026: /* Baseplate Preparation */</p>
<hr />
<div>The following is a basic outline of our procedure for implanting a GRIN lens above the hippocampus for CA1 imaging. For videos of surgery and baseplating, see the [[Online Workshop]]. Also, see the recent Nature Protocols paper from the Stuber Lab ([http://www.nature.com/nprot/journal/v11/n3/full/nprot.2016.021.html link]) that has a very detailed description of the surgery. <br />
<br />
In order to image, animals will need to undergo three surgical procedures: a virus injection, a GRIN lens implantation, and baseplate attachment.<br />
<br />
== Virus Injection ==<br />
Before implanting the GRIN lens, you will need to inject a fluorescent indicator such as GCaMP6. We have generally used AAV1.Syn.GCaMP6f.WPRE.SV40 from Penn Vector (Item number AV-1-PV2822) and had great success in dorsal CA1. You may also be able to use GCaMP6 transgenic mice, but we have not tested whether any of the newer lines are bright enough to be imaged.<br />
<br />
== GRIN Lens Implantation==<br />
<br />
=== Basic Equipment Needed ===<br />
You will only need basic surgical equipment to perform GRIN lens implantation surgeries. This includes a mouse stereotax, an isoflurane vaporizor, surgical heat pad, stereo surgical microscope, light source, dental drill, and bead sterilizer. For recommended equipment, see the master [https://docs.google.com/spreadsheets/d/12H71DU2QX8d7efUE4yNuikBEiIzKaXjYqdc0A-oLNSw/edit?usp=sharing parts list].<br />
<br />
You will need the following basic tools and supplies: fine forceps, blunt forceps, fine scissors, scalpel and blade, small skull screws, drill bits, cyanoacrylate glue, dental cement, Kwik-Sil, and [[cortex buffer]]<br />
<br />
=== Aspirator and Lens Holder ===<br />
You will need an aspirator to remove cortex above the hippocampus. This can be built very simply with a vacuum line (or pump), a liquid trap, 1 ml syringe (with a hole to control suction), blunt needles, tubing and connectors, and hot glue.<br />
[[File:Aspirator.PNG|center|500px]]<br />
<br />
You can easily build a GRIN lens holder by using a suction system to hold the lens in place as it is dropped into the brain for implantation. To build, start with two 1ml micropipette tips. Cut the first so that the tip is just larger than the diameter of the GRIN lens you will be using. Cut the second tip so that the end is just smaller than the size of the GRIN lens. Connect this end to a 1ml syringe and connect to a vacuum line. This will then hold the lens in place during implantation. You can then use tape to attach this holder to a stereotax. Another solution we have used is a basic drill chuck, but this is only good for large (2mm) lenses.<br />
[[File:LensHolder.PNG|center|500px]]<br />
<br />
=== Skull Preparation ===<br />
You must first prepare the skull for implantation. We recommend shaving the head area and sterilizing with 3 alternating betadine and ethanol scrubs. Next, remove the scalp with scissors and clean the skull with hydrogen peroxide and saline. Scrape and score the skull to increase the bond with the glue. We also detach the neck muscle from the skull in order to reduce pull on the skull and prevent muscle growth that can make the implant less stable. Finally we insert a skull screw on the opposite side of the skull to enhance the stability of the implant.<br />
<br />
=== Craniotomy and Aspiration ===<br />
Using a stereotax, align the skull and find the location for the implant. Using a drill, make 4 guide holes to outline the lens placement. For CA1, we recommend offsetting the lens 0.5mm to the medial side of the virus injection to prevent imaging any damage induced by the virus injection. Connect the guide holes to create a circular craniotomy and peel off the circular skull fragment. Do not worry about blood as the top layers of cortex will be removed by aspiration. Make absolutely sure that the craniotomy is at least the size of your GRIN lens! Cover with cortex buffer and scrape around the outside of the craniotomy to remove any excess bone or dura. <br />
<br />
While keeping a constant flow of cortex buffer use the aspirator to slowly remove the cortex above the hippocampus. Continue removing cortex until you reach the white striations of the corpus callosum. Very slowly and carefully remove the horizontal white striations of the corpus callosum until you reach the striations that go vertical. When you reach these striations, stop aspirating and clean up the sides so that the lens can be placed into the brain. Continue washing with cortex buffer until all bleeding stops and keep the brain constantly flushed with cortex buffer.<br />
<br />
=== Lens implantation ===<br />
Attach GRIN lens to lens holder on stereotax. Align the lens with the top of the skull at the most posterior part of the craniotomy and quickly insert the lens 1.35 mm below the top of the skull. Remove any excess liquid with the aspirator. Connect the outside of the GRIN lens with the skull and skull screw using cyanoacrylate glue and let fully dry or use a glue accelerator. Remove the lens holder by simply removing the vacuum suction (you can just pinch the tubing) and withdrawing the holder. Next, cover the entire skull with cyanoacrylate glue, and then cover with dental cement. Let dry completely and cover with Kwik-Sil to protect the lens. Remove from stereotax and allow the animal to recover. Give the animal amoxicillin (or equivalent) for 7 days, and daily injections of carprofen and dexamethasone for 7 days.<br />
<br />
=== Important tips to remember ===<br />
*Try to keep the surgery as sterile as possible. Infections will destroy imaging and the ability to keep the same cells over many days.<br />
<br />
*Try to keep isoflurane as low as possible. Surgeries can take 2-6 hours depending on experience.<br />
<br />
*Aspirations will take a lot of practice. Go slow at first.<br />
<br />
*Depth will be trial and error. If the first few animals don't give good images, check the depth using histology and adjust accordingly.<br />
<br />
<br />
Please feel free to add any tips that you have!<br />
<br />
== Baseplating Protocol ==<br />
<br />
<br />
=== Basic Equipment needed ===<br />
You will only need basic surgical equipment to baseplate. This includes a mouse stereotax, an isoflurane vaporizer, surgical heat pad, stereo surgical microscope, light source, and a dental drill. <br />
<br />
You will also need cyanoacrylate glue, fine forceps, double distilled water, lens paper, and dental cement<br />
=== Baseplate Preparation ===<br />
Before affixing the baseplate on an animal, you will need to prepare the baseplate. <br />
<br />
The first step is to insert magnets into the appropriate holes on the baseplate. To ensure that the polarity of the magnets is correct throughout the baseplate preparation, use a reference Miniscope to hold onto the magnets. Mark the top magnet with a sharpie, as seen in the picture below. Next, put the top magnet on the head of a screwdriver with the marked side facing away from the screwdriver. Apply cyanoacrylate glue on the edge of the magnet, then push the magnet through the appropriate hole. Ensure that the magnet goes through the hole completely and the top side of the baseplate is flat, as the quality of imaging relies on the Miniscope sitting flush on the baseplate. Repeat this procedure for the next 2 magnets. Finally, put a coat of cyanoacrylate glue on the bottom side of the baseplate and leave the baseplate to dry.<br />
<br />
Once the baseplate has dried, score the bottom and the sides of the baseplate using a dental drill to allow the dental glue to adhere the baseplate and skull well.<br />
<br />
=== Cap Preparation ===<br />
Similar to the baseplate preparation, use a reference Miniscope to hold onto the magnets. Mark the top magnet with a sharpie, and put the magnet on the head of a screwdriver with the marked side facing toward the screwdriver. Coat the magnet with cyanoacrylate glue and push the magnet into the appropriate hole. Repeat for the remaining two magnets.<br />
=== Baseplating ===<br />
Connect a Miniscope to a DAQ and initialize the Miniscope software on a computer. Attach your prepared baseplate onto the bottom of the Miniscope using a set screw.<br />
<br />
Using a stereotax and isoflurane vaporizer, anesthetize the animal and fix ear bars until the animal is stable. Carefully remove the Kwik-Sil off the top of the animal’s GRIN lens with blunt forceps being sure not to scratch the lens. Under a stereo surgical microscope, check to see if there is debris on the top of the GRIN lens. If there is, dampen a sheet of lens paper using double distilled water. Wipe the top of the GRIN lens using fine forceps and lens paper, being sure to only wipe laterally and not to apply direct downward pressure.<br />
<br />
After the top of the GRIN lens is clean, explore the field of view with the Miniscope. Adjust the LED brightness and focus accordingly. We advise that you use maximum gain and low LED excitation as to not photobleach the brain during baseplating. After finding your ideal view, practice taking the Miniscope off and on and finding the same view. <br />
<br />
When you feel comfortable repeatedly finding the view with the Miniscope, apply dental cement in a “C” pattern around the GRIN lens, making sure not to touch the lens itself. It will be easier if your first application of dental cement is more viscous. Put the Miniscope on top of the lens and find your ideal view. Hold the Miniscope steadily as the first application of dental cement dries, making sure that your view does not change. <br />
After the first application of dental cement dries, mix more dental cement and apply it on the posterior edge of the baseplate, making sure it seals to the skull. At this point, you can also build up dental cement along the wall of the baseplate on the animal’s right side. Be sure to avoid applying dental cement onto the camera itself and adhering it to the baseplate. Once the dental cement dries and you feel confident that the baseplate will no longer shift, carefully remove the Miniscope from the baseplate by unscrewing the set screw and lifting it off. <br />
<br />
Apply dental cement to the anterior edge of the baseplate and fill in the gaps between the baseplate and skull on the animal’s left side. Allow the dental cement to dry and attach the plastic cap onto the top of the baseplate with a set screw.</div>Chrisl1026http://miniscope.org/index.php?title=Surgery_Protocol&diff=1516Surgery Protocol2016-09-21T03:42:00Z<p>Chrisl1026: /* Baseplate Preparation */</p>
<hr />
<div>The following is a basic outline of our procedure for implanting a GRIN lens above the hippocampus for CA1 imaging. For videos of surgery and baseplating, see the [[Online Workshop]]. Also, see the recent Nature Protocols paper from the Stuber Lab ([http://www.nature.com/nprot/journal/v11/n3/full/nprot.2016.021.html link]) that has a very detailed description of the surgery. <br />
<br />
In order to image, animals will need to undergo three surgical procedures: a virus injection, a GRIN lens implantation, and baseplate attachment.<br />
<br />
== Virus Injection ==<br />
Before implanting the GRIN lens, you will need to inject a fluorescent indicator such as GCaMP6. We have generally used AAV1.Syn.GCaMP6f.WPRE.SV40 from Penn Vector (Item number AV-1-PV2822) and had great success in dorsal CA1. You may also be able to use GCaMP6 transgenic mice, but we have not tested whether any of the newer lines are bright enough to be imaged.<br />
<br />
== GRIN Lens Implantation==<br />
<br />
=== Basic Equipment Needed ===<br />
You will only need basic surgical equipment to perform GRIN lens implantation surgeries. This includes a mouse stereotax, an isoflurane vaporizor, surgical heat pad, stereo surgical microscope, light source, dental drill, and bead sterilizer. For recommended equipment, see the master [https://docs.google.com/spreadsheets/d/12H71DU2QX8d7efUE4yNuikBEiIzKaXjYqdc0A-oLNSw/edit?usp=sharing parts list].<br />
<br />
You will need the following basic tools and supplies: fine forceps, blunt forceps, fine scissors, scalpel and blade, small skull screws, drill bits, cyanoacrylate glue, dental cement, Kwik-Sil, and [[cortex buffer]]<br />
<br />
=== Aspirator and Lens Holder ===<br />
You will need an aspirator to remove cortex above the hippocampus. This can be built very simply with a vacuum line (or pump), a liquid trap, 1 ml syringe (with a hole to control suction), blunt needles, tubing and connectors, and hot glue.<br />
[[File:Aspirator.PNG|center|500px]]<br />
<br />
You can easily build a GRIN lens holder by using a suction system to hold the lens in place as it is dropped into the brain for implantation. To build, start with two 1ml micropipette tips. Cut the first so that the tip is just larger than the diameter of the GRIN lens you will be using. Cut the second tip so that the end is just smaller than the size of the GRIN lens. Connect this end to a 1ml syringe and connect to a vacuum line. This will then hold the lens in place during implantation. You can then use tape to attach this holder to a stereotax. Another solution we have used is a basic drill chuck, but this is only good for large (2mm) lenses.<br />
[[File:LensHolder.PNG|center|500px]]<br />
<br />
=== Skull Preparation ===<br />
You must first prepare the skull for implantation. We recommend shaving the head area and sterilizing with 3 alternating betadine and ethanol scrubs. Next, remove the scalp with scissors and clean the skull with hydrogen peroxide and saline. Scrape and score the skull to increase the bond with the glue. We also detach the neck muscle from the skull in order to reduce pull on the skull and prevent muscle growth that can make the implant less stable. Finally we insert a skull screw on the opposite side of the skull to enhance the stability of the implant.<br />
<br />
=== Craniotomy and Aspiration ===<br />
Using a stereotax, align the skull and find the location for the implant. Using a drill, make 4 guide holes to outline the lens placement. For CA1, we recommend offsetting the lens 0.5mm to the medial side of the virus injection to prevent imaging any damage induced by the virus injection. Connect the guide holes to create a circular craniotomy and peel off the circular skull fragment. Do not worry about blood as the top layers of cortex will be removed by aspiration. Make absolutely sure that the craniotomy is at least the size of your GRIN lens! Cover with cortex buffer and scrape around the outside of the craniotomy to remove any excess bone or dura. <br />
<br />
While keeping a constant flow of cortex buffer use the aspirator to slowly remove the cortex above the hippocampus. Continue removing cortex until you reach the white striations of the corpus callosum. Very slowly and carefully remove the horizontal white striations of the corpus callosum until you reach the striations that go vertical. When you reach these striations, stop aspirating and clean up the sides so that the lens can be placed into the brain. Continue washing with cortex buffer until all bleeding stops and keep the brain constantly flushed with cortex buffer.<br />
<br />
=== Lens implantation ===<br />
Attach GRIN lens to lens holder on stereotax. Align the lens with the top of the skull at the most posterior part of the craniotomy and quickly insert the lens 1.35 mm below the top of the skull. Remove any excess liquid with the aspirator. Connect the outside of the GRIN lens with the skull and skull screw using cyanoacrylate glue and let fully dry or use a glue accelerator. Remove the lens holder by simply removing the vacuum suction (you can just pinch the tubing) and withdrawing the holder. Next, cover the entire skull with cyanoacrylate glue, and then cover with dental cement. Let dry completely and cover with Kwik-Sil to protect the lens. Remove from stereotax and allow the animal to recover. Give the animal amoxicillin (or equivalent) for 7 days, and daily injections of carprofen and dexamethasone for 7 days.<br />
<br />
=== Important tips to remember ===<br />
*Try to keep the surgery as sterile as possible. Infections will destroy imaging and the ability to keep the same cells over many days.<br />
<br />
*Try to keep isoflurane as low as possible. Surgeries can take 2-6 hours depending on experience.<br />
<br />
*Aspirations will take a lot of practice. Go slow at first.<br />
<br />
*Depth will be trial and error. If the first few animals don't give good images, check the depth using histology and adjust accordingly.<br />
<br />
<br />
Please feel free to add any tips that you have!<br />
<br />
== Baseplating Protocol ==<br />
<br />
<br />
=== Basic Equipment needed ===<br />
You will only need basic surgical equipment to baseplate. This includes a mouse stereotax, an isoflurane vaporizer, surgical heat pad, stereo surgical microscope, light source, and a dental drill. <br />
<br />
You will also need cyanoacrylate glue, fine forceps, double distilled water, lens paper, and dental cement<br />
=== Baseplate Preparation ===<br />
Before affixing the baseplate on an animal, you will need to prepare the baseplate. <br />
<br />
The first step is to insert magnets into the appropriate holes on the baseplate. To ensure that the polarity of the magnets is correct throughout the baseplate preparation, use a reference Miniscope to hold onto the magnets. Mark the top magnet with a sharpie, as seen in the picture below. Next, put the top magnet on the head of a screwdriver with the marked side facing away from the screwdriver. Apply cyanoacrylate glue on the edge of the magnet, then push the magnet through the appropriate hole. Ensure that the magnet goes through the hole completely and the top side of the baseplate is flat, as the quality of imaging relies on the Miniscope sitting flush on the baseplate. Repeat this procedure for the next 2 magnets. Finally, put a coat of cyanoacrylate glue on the bottom side of the baseplate and leave the baseplate to dry.<br />
<br />
Once the baseplate has dried, score the bottom and the sides of the baseplate using a dental drill to allow the dental glue to adhere the baseplate and skull well.<br />
[[File:Image 1062.jpg|center|200px]]<br />
<br />
=== Cap Preparation ===<br />
Similar to the baseplate preparation, use a reference Miniscope to hold onto the magnets. Mark the top magnet with a sharpie, and put the magnet on the head of a screwdriver with the marked side facing toward the screwdriver. Coat the magnet with cyanoacrylate glue and push the magnet into the appropriate hole. Repeat for the remaining two magnets.<br />
=== Baseplating ===<br />
Connect a Miniscope to a DAQ and initialize the Miniscope software on a computer. Attach your prepared baseplate onto the bottom of the Miniscope using a set screw.<br />
<br />
Using a stereotax and isoflurane vaporizer, anesthetize the animal and fix ear bars until the animal is stable. Carefully remove the Kwik-Sil off the top of the animal’s GRIN lens with blunt forceps being sure not to scratch the lens. Under a stereo surgical microscope, check to see if there is debris on the top of the GRIN lens. If there is, dampen a sheet of lens paper using double distilled water. Wipe the top of the GRIN lens using fine forceps and lens paper, being sure to only wipe laterally and not to apply direct downward pressure.<br />
<br />
After the top of the GRIN lens is clean, explore the field of view with the Miniscope. Adjust the LED brightness and focus accordingly. We advise that you use maximum gain and low LED excitation as to not photobleach the brain during baseplating. After finding your ideal view, practice taking the Miniscope off and on and finding the same view. <br />
<br />
When you feel comfortable repeatedly finding the view with the Miniscope, apply dental cement in a “C” pattern around the GRIN lens, making sure not to touch the lens itself. It will be easier if your first application of dental cement is more viscous. Put the Miniscope on top of the lens and find your ideal view. Hold the Miniscope steadily as the first application of dental cement dries, making sure that your view does not change. <br />
After the first application of dental cement dries, mix more dental cement and apply it on the posterior edge of the baseplate, making sure it seals to the skull. At this point, you can also build up dental cement along the wall of the baseplate on the animal’s right side. Be sure to avoid applying dental cement onto the camera itself and adhering it to the baseplate. Once the dental cement dries and you feel confident that the baseplate will no longer shift, carefully remove the Miniscope from the baseplate by unscrewing the set screw and lifting it off. <br />
<br />
Apply dental cement to the anterior edge of the baseplate and fill in the gaps between the baseplate and skull on the animal’s left side. Allow the dental cement to dry and attach the plastic cap onto the top of the baseplate with a set screw.</div>Chrisl1026http://miniscope.org/index.php?title=Surgery_Protocol&diff=1514Surgery Protocol2016-09-21T02:22:02Z<p>Chrisl1026: /* Baseplate Preparation */</p>
<hr />
<div>The following is a basic outline of our procedure for implanting a GRIN lens above the hippocampus for CA1 imaging. For videos of surgery and baseplating, see the [[Online Workshop]]. Also, see the recent Nature Protocols paper from the Stuber Lab ([http://www.nature.com/nprot/journal/v11/n3/full/nprot.2016.021.html link]) that has a very detailed description of the surgery. <br />
<br />
In order to image, animals will need to undergo three surgical procedures: a virus injection, a GRIN lens implantation, and baseplate attachment.<br />
<br />
== Virus Injection ==<br />
Before implanting the GRIN lens, you will need to inject a fluorescent indicator such as GCaMP6. We have generally used AAV1.Syn.GCaMP6f.WPRE.SV40 from Penn Vector (Item number AV-1-PV2822) and had great success in dorsal CA1. You may also be able to use GCaMP6 transgenic mice, but we have not tested whether any of the newer lines are bright enough to be imaged.<br />
<br />
== GRIN Lens Implantation==<br />
<br />
=== Basic Equipment Needed ===<br />
You will only need basic surgical equipment to perform GRIN lens implantation surgeries. This includes a mouse stereotax, an isoflurane vaporizor, surgical heat pad, stereo surgical microscope, light source, dental drill, and bead sterilizer. For recommended equipment, see the master [https://docs.google.com/spreadsheets/d/12H71DU2QX8d7efUE4yNuikBEiIzKaXjYqdc0A-oLNSw/edit?usp=sharing parts list].<br />
<br />
You will need the following basic tools and supplies: fine forceps, blunt forceps, fine scissors, scalpel and blade, small skull screws, drill bits, cyanoacrylate glue, dental cement, Kwik-Sil, and [[cortex buffer]]<br />
<br />
=== Aspirator and Lens Holder ===<br />
You will need an aspirator to remove cortex above the hippocampus. This can be built very simply with a vacuum line (or pump), a liquid trap, 1 ml syringe (with a hole to control suction), blunt needles, tubing and connectors, and hot glue.<br />
[[File:Aspirator.PNG|center|500px]]<br />
<br />
You can easily build a GRIN lens holder by using a suction system to hold the lens in place as it is dropped into the brain for implantation. To build, start with two 1ml micropipette tips. Cut the first so that the tip is just larger than the diameter of the GRIN lens you will be using. Cut the second tip so that the end is just smaller than the size of the GRIN lens. Connect this end to a 1ml syringe and connect to a vacuum line. This will then hold the lens in place during implantation. You can then use tape to attach this holder to a stereotax. Another solution we have used is a basic drill chuck, but this is only good for large (2mm) lenses.<br />
[[File:LensHolder.PNG|center|500px]]<br />
<br />
=== Skull Preparation ===<br />
You must first prepare the skull for implantation. We recommend shaving the head area and sterilizing with 3 alternating betadine and ethanol scrubs. Next, remove the scalp with scissors and clean the skull with hydrogen peroxide and saline. Scrape and score the skull to increase the bond with the glue. We also detach the neck muscle from the skull in order to reduce pull on the skull and prevent muscle growth that can make the implant less stable. Finally we insert a skull screw on the opposite side of the skull to enhance the stability of the implant.<br />
<br />
=== Craniotomy and Aspiration ===<br />
Using a stereotax, align the skull and find the location for the implant. Using a drill, make 4 guide holes to outline the lens placement. For CA1, we recommend offsetting the lens 0.5mm to the medial side of the virus injection to prevent imaging any damage induced by the virus injection. Connect the guide holes to create a circular craniotomy and peel off the circular skull fragment. Do not worry about blood as the top layers of cortex will be removed by aspiration. Make absolutely sure that the craniotomy is at least the size of your GRIN lens! Cover with cortex buffer and scrape around the outside of the craniotomy to remove any excess bone or dura. <br />
<br />
While keeping a constant flow of cortex buffer use the aspirator to slowly remove the cortex above the hippocampus. Continue removing cortex until you reach the white striations of the corpus callosum. Very slowly and carefully remove the horizontal white striations of the corpus callosum until you reach the striations that go vertical. When you reach these striations, stop aspirating and clean up the sides so that the lens can be placed into the brain. Continue washing with cortex buffer until all bleeding stops and keep the brain constantly flushed with cortex buffer.<br />
<br />
=== Lens implantation ===<br />
Attach GRIN lens to lens holder on stereotax. Align the lens with the top of the skull at the most posterior part of the craniotomy and quickly insert the lens 1.35 mm below the top of the skull. Remove any excess liquid with the aspirator. Connect the outside of the GRIN lens with the skull and skull screw using cyanoacrylate glue and let fully dry or use a glue accelerator. Remove the lens holder by simply removing the vacuum suction (you can just pinch the tubing) and withdrawing the holder. Next, cover the entire skull with cyanoacrylate glue, and then cover with dental cement. Let dry completely and cover with Kwik-Sil to protect the lens. Remove from stereotax and allow the animal to recover. Give the animal amoxicillin (or equivalent) for 7 days, and daily injections of carprofen and dexamethasone for 7 days.<br />
<br />
=== Important tips to remember ===<br />
*Try to keep the surgery as sterile as possible. Infections will destroy imaging and the ability to keep the same cells over many days.<br />
<br />
*Try to keep isoflurane as low as possible. Surgeries can take 2-6 hours depending on experience.<br />
<br />
*Aspirations will take a lot of practice. Go slow at first.<br />
<br />
*Depth will be trial and error. If the first few animals don't give good images, check the depth using histology and adjust accordingly.<br />
<br />
<br />
Please feel free to add any tips that you have!<br />
<br />
== Baseplating Protocol ==<br />
<br />
<br />
=== Basic Equipment needed ===<br />
You will only need basic surgical equipment to baseplate. This includes a mouse stereotax, an isoflurane vaporizer, surgical heat pad, stereo surgical microscope, light source, and a dental drill. <br />
<br />
You will also need cyanoacrylate glue, fine forceps, double distilled water, lens paper, and dental cement<br />
=== Baseplate Preparation ===<br />
Before affixing the baseplate on an animal, you will need to prepare the baseplate. <br />
<br />
The first step is to insert magnets into the appropriate holes on the baseplate. To ensure that the polarity of the magnets is correct throughout the baseplate preparation, use a reference Miniscope to hold onto the magnets. Mark the top magnet with a sharpie, as seen in the picture below. Next, put the top magnet on the head of a screwdriver with the marked side facing away from the screwdriver. Apply cyanoacrylate glue on the edge of the magnet, then push the magnet through the appropriate hole. Ensure that the magnet goes through the hole completely and the top side of the baseplate is flat, as the quality of imaging relies on the Miniscope sitting flush on the baseplate. Repeat this procedure for the next 2 magnets. Finally, put a coat of cyanoacrylate glue on the bottom side of the baseplate and leave the baseplate to dry.<br />
<br />
Once the baseplate has dried, score the bottom and the sides of the baseplate using a dental drill to allow the dental glue to adhere the baseplate and skull well.<br />
[[File:BP1.jpg|center|100px]]<br />
<br />
=== Cap Preparation ===<br />
Similar to the baseplate preparation, use a reference Miniscope to hold onto the magnets. Mark the top magnet with a sharpie, and put the magnet on the head of a screwdriver with the marked side facing toward the screwdriver. Coat the magnet with cyanoacrylate glue and push the magnet into the appropriate hole. Repeat for the remaining two magnets.<br />
=== Baseplating ===<br />
Connect a Miniscope to a DAQ and initialize the Miniscope software on a computer. Attach your prepared baseplate onto the bottom of the Miniscope using a set screw.<br />
<br />
Using a stereotax and isoflurane vaporizer, anesthetize the animal and fix ear bars until the animal is stable. Carefully remove the Kwik-Sil off the top of the animal’s GRIN lens with blunt forceps being sure not to scratch the lens. Under a stereo surgical microscope, check to see if there is debris on the top of the GRIN lens. If there is, dampen a sheet of lens paper using double distilled water. Wipe the top of the GRIN lens using fine forceps and lens paper, being sure to only wipe laterally and not to apply direct downward pressure.<br />
<br />
After the top of the GRIN lens is clean, explore the field of view with the Miniscope. Adjust the LED brightness and focus accordingly. We advise that you use maximum gain and low LED excitation as to not photobleach the brain during baseplating. After finding your ideal view, practice taking the Miniscope off and on and finding the same view. <br />
<br />
When you feel comfortable repeatedly finding the view with the Miniscope, apply dental cement in a “C” pattern around the GRIN lens, making sure not to touch the lens itself. It will be easier if your first application of dental cement is more viscous. Put the Miniscope on top of the lens and find your ideal view. Hold the Miniscope steadily as the first application of dental cement dries, making sure that your view does not change. <br />
After the first application of dental cement dries, mix more dental cement and apply it on the posterior edge of the baseplate, making sure it seals to the skull. At this point, you can also build up dental cement along the wall of the baseplate on the animal’s right side. Be sure to avoid applying dental cement onto the camera itself and adhering it to the baseplate. Once the dental cement dries and you feel confident that the baseplate will no longer shift, carefully remove the Miniscope from the baseplate by unscrewing the set screw and lifting it off. <br />
<br />
Apply dental cement to the anterior edge of the baseplate and fill in the gaps between the baseplate and skull on the animal’s left side. Allow the dental cement to dry and attach the plastic cap onto the top of the baseplate with a set screw.</div>Chrisl1026http://miniscope.org/index.php?title=Surgery_Protocol&diff=1512Surgery Protocol2016-09-21T02:19:32Z<p>Chrisl1026: /* Baseplate Preparation */</p>
<hr />
<div>The following is a basic outline of our procedure for implanting a GRIN lens above the hippocampus for CA1 imaging. For videos of surgery and baseplating, see the [[Online Workshop]]. Also, see the recent Nature Protocols paper from the Stuber Lab ([http://www.nature.com/nprot/journal/v11/n3/full/nprot.2016.021.html link]) that has a very detailed description of the surgery. <br />
<br />
In order to image, animals will need to undergo three surgical procedures: a virus injection, a GRIN lens implantation, and baseplate attachment.<br />
<br />
== Virus Injection ==<br />
Before implanting the GRIN lens, you will need to inject a fluorescent indicator such as GCaMP6. We have generally used AAV1.Syn.GCaMP6f.WPRE.SV40 from Penn Vector (Item number AV-1-PV2822) and had great success in dorsal CA1. You may also be able to use GCaMP6 transgenic mice, but we have not tested whether any of the newer lines are bright enough to be imaged.<br />
<br />
== GRIN Lens Implantation==<br />
<br />
=== Basic Equipment Needed ===<br />
You will only need basic surgical equipment to perform GRIN lens implantation surgeries. This includes a mouse stereotax, an isoflurane vaporizor, surgical heat pad, stereo surgical microscope, light source, dental drill, and bead sterilizer. For recommended equipment, see the master [https://docs.google.com/spreadsheets/d/12H71DU2QX8d7efUE4yNuikBEiIzKaXjYqdc0A-oLNSw/edit?usp=sharing parts list].<br />
<br />
You will need the following basic tools and supplies: fine forceps, blunt forceps, fine scissors, scalpel and blade, small skull screws, drill bits, cyanoacrylate glue, dental cement, Kwik-Sil, and [[cortex buffer]]<br />
<br />
=== Aspirator and Lens Holder ===<br />
You will need an aspirator to remove cortex above the hippocampus. This can be built very simply with a vacuum line (or pump), a liquid trap, 1 ml syringe (with a hole to control suction), blunt needles, tubing and connectors, and hot glue.<br />
[[File:Aspirator.PNG|center|500px]]<br />
<br />
You can easily build a GRIN lens holder by using a suction system to hold the lens in place as it is dropped into the brain for implantation. To build, start with two 1ml micropipette tips. Cut the first so that the tip is just larger than the diameter of the GRIN lens you will be using. Cut the second tip so that the end is just smaller than the size of the GRIN lens. Connect this end to a 1ml syringe and connect to a vacuum line. This will then hold the lens in place during implantation. You can then use tape to attach this holder to a stereotax. Another solution we have used is a basic drill chuck, but this is only good for large (2mm) lenses.<br />
[[File:LensHolder.PNG|center|500px]]<br />
<br />
=== Skull Preparation ===<br />
You must first prepare the skull for implantation. We recommend shaving the head area and sterilizing with 3 alternating betadine and ethanol scrubs. Next, remove the scalp with scissors and clean the skull with hydrogen peroxide and saline. Scrape and score the skull to increase the bond with the glue. We also detach the neck muscle from the skull in order to reduce pull on the skull and prevent muscle growth that can make the implant less stable. Finally we insert a skull screw on the opposite side of the skull to enhance the stability of the implant.<br />
<br />
=== Craniotomy and Aspiration ===<br />
Using a stereotax, align the skull and find the location for the implant. Using a drill, make 4 guide holes to outline the lens placement. For CA1, we recommend offsetting the lens 0.5mm to the medial side of the virus injection to prevent imaging any damage induced by the virus injection. Connect the guide holes to create a circular craniotomy and peel off the circular skull fragment. Do not worry about blood as the top layers of cortex will be removed by aspiration. Make absolutely sure that the craniotomy is at least the size of your GRIN lens! Cover with cortex buffer and scrape around the outside of the craniotomy to remove any excess bone or dura. <br />
<br />
While keeping a constant flow of cortex buffer use the aspirator to slowly remove the cortex above the hippocampus. Continue removing cortex until you reach the white striations of the corpus callosum. Very slowly and carefully remove the horizontal white striations of the corpus callosum until you reach the striations that go vertical. When you reach these striations, stop aspirating and clean up the sides so that the lens can be placed into the brain. Continue washing with cortex buffer until all bleeding stops and keep the brain constantly flushed with cortex buffer.<br />
<br />
=== Lens implantation ===<br />
Attach GRIN lens to lens holder on stereotax. Align the lens with the top of the skull at the most posterior part of the craniotomy and quickly insert the lens 1.35 mm below the top of the skull. Remove any excess liquid with the aspirator. Connect the outside of the GRIN lens with the skull and skull screw using cyanoacrylate glue and let fully dry or use a glue accelerator. Remove the lens holder by simply removing the vacuum suction (you can just pinch the tubing) and withdrawing the holder. Next, cover the entire skull with cyanoacrylate glue, and then cover with dental cement. Let dry completely and cover with Kwik-Sil to protect the lens. Remove from stereotax and allow the animal to recover. Give the animal amoxicillin (or equivalent) for 7 days, and daily injections of carprofen and dexamethasone for 7 days.<br />
<br />
=== Important tips to remember ===<br />
*Try to keep the surgery as sterile as possible. Infections will destroy imaging and the ability to keep the same cells over many days.<br />
<br />
*Try to keep isoflurane as low as possible. Surgeries can take 2-6 hours depending on experience.<br />
<br />
*Aspirations will take a lot of practice. Go slow at first.<br />
<br />
*Depth will be trial and error. If the first few animals don't give good images, check the depth using histology and adjust accordingly.<br />
<br />
<br />
Please feel free to add any tips that you have!<br />
<br />
== Baseplating Protocol ==<br />
<br />
<br />
=== Basic Equipment needed ===<br />
You will only need basic surgical equipment to baseplate. This includes a mouse stereotax, an isoflurane vaporizer, surgical heat pad, stereo surgical microscope, light source, and a dental drill. <br />
<br />
You will also need cyanoacrylate glue, fine forceps, double distilled water, lens paper, and dental cement<br />
=== Baseplate Preparation ===<br />
Before affixing the baseplate on an animal, you will need to prepare the baseplate. <br />
<br />
The first step is to insert magnets into the appropriate holes on the baseplate. To ensure that the polarity of the magnets is correct throughout the baseplate preparation, use a reference Miniscope to hold onto the magnets. Mark the top magnet with a sharpie, as seen in the picture below. Next, put the top magnet on the head of a screwdriver with the marked side facing away from the screwdriver. Apply cyanoacrylate glue on the edge of the magnet, then push the magnet through the appropriate hole. Ensure that the magnet goes through the hole completely and the top side of the baseplate is flat, as the quality of imaging relies on the Miniscope sitting flush on the baseplate. Repeat this procedure for the next 2 magnets. Finally, put a coat of cyanoacrylate glue on the bottom side of the baseplate and leave the baseplate to dry.<br />
<br />
Once the baseplate has dried, score the bottom and the sides of the baseplate using a dental drill to allow the dental glue to adhere the baseplate and skull well.<br />
[[File:BP1.jpg]]<br />
<br />
=== Cap Preparation ===<br />
Similar to the baseplate preparation, use a reference Miniscope to hold onto the magnets. Mark the top magnet with a sharpie, and put the magnet on the head of a screwdriver with the marked side facing toward the screwdriver. Coat the magnet with cyanoacrylate glue and push the magnet into the appropriate hole. Repeat for the remaining two magnets.<br />
=== Baseplating ===<br />
Connect a Miniscope to a DAQ and initialize the Miniscope software on a computer. Attach your prepared baseplate onto the bottom of the Miniscope using a set screw.<br />
<br />
Using a stereotax and isoflurane vaporizer, anesthetize the animal and fix ear bars until the animal is stable. Carefully remove the Kwik-Sil off the top of the animal’s GRIN lens with blunt forceps being sure not to scratch the lens. Under a stereo surgical microscope, check to see if there is debris on the top of the GRIN lens. If there is, dampen a sheet of lens paper using double distilled water. Wipe the top of the GRIN lens using fine forceps and lens paper, being sure to only wipe laterally and not to apply direct downward pressure.<br />
<br />
After the top of the GRIN lens is clean, explore the field of view with the Miniscope. Adjust the LED brightness and focus accordingly. We advise that you use maximum gain and low LED excitation as to not photobleach the brain during baseplating. After finding your ideal view, practice taking the Miniscope off and on and finding the same view. <br />
<br />
When you feel comfortable repeatedly finding the view with the Miniscope, apply dental cement in a “C” pattern around the GRIN lens, making sure not to touch the lens itself. It will be easier if your first application of dental cement is more viscous. Put the Miniscope on top of the lens and find your ideal view. Hold the Miniscope steadily as the first application of dental cement dries, making sure that your view does not change. <br />
After the first application of dental cement dries, mix more dental cement and apply it on the posterior edge of the baseplate, making sure it seals to the skull. At this point, you can also build up dental cement along the wall of the baseplate on the animal’s right side. Be sure to avoid applying dental cement onto the camera itself and adhering it to the baseplate. Once the dental cement dries and you feel confident that the baseplate will no longer shift, carefully remove the Miniscope from the baseplate by unscrewing the set screw and lifting it off. <br />
<br />
Apply dental cement to the anterior edge of the baseplate and fill in the gaps between the baseplate and skull on the animal’s left side. Allow the dental cement to dry and attach the plastic cap onto the top of the baseplate with a set screw.</div>Chrisl1026http://miniscope.org/index.php?title=Surgery_Protocol&diff=1510Surgery Protocol2016-09-21T02:15:29Z<p>Chrisl1026: /* Baseplate Preparation */</p>
<hr />
<div>The following is a basic outline of our procedure for implanting a GRIN lens above the hippocampus for CA1 imaging. For videos of surgery and baseplating, see the [[Online Workshop]]. Also, see the recent Nature Protocols paper from the Stuber Lab ([http://www.nature.com/nprot/journal/v11/n3/full/nprot.2016.021.html link]) that has a very detailed description of the surgery. <br />
<br />
In order to image, animals will need to undergo three surgical procedures: a virus injection, a GRIN lens implantation, and baseplate attachment.<br />
<br />
== Virus Injection ==<br />
Before implanting the GRIN lens, you will need to inject a fluorescent indicator such as GCaMP6. We have generally used AAV1.Syn.GCaMP6f.WPRE.SV40 from Penn Vector (Item number AV-1-PV2822) and had great success in dorsal CA1. You may also be able to use GCaMP6 transgenic mice, but we have not tested whether any of the newer lines are bright enough to be imaged.<br />
<br />
== GRIN Lens Implantation==<br />
<br />
=== Basic Equipment Needed ===<br />
You will only need basic surgical equipment to perform GRIN lens implantation surgeries. This includes a mouse stereotax, an isoflurane vaporizor, surgical heat pad, stereo surgical microscope, light source, dental drill, and bead sterilizer. For recommended equipment, see the master [https://docs.google.com/spreadsheets/d/12H71DU2QX8d7efUE4yNuikBEiIzKaXjYqdc0A-oLNSw/edit?usp=sharing parts list].<br />
<br />
You will need the following basic tools and supplies: fine forceps, blunt forceps, fine scissors, scalpel and blade, small skull screws, drill bits, cyanoacrylate glue, dental cement, Kwik-Sil, and [[cortex buffer]]<br />
<br />
=== Aspirator and Lens Holder ===<br />
You will need an aspirator to remove cortex above the hippocampus. This can be built very simply with a vacuum line (or pump), a liquid trap, 1 ml syringe (with a hole to control suction), blunt needles, tubing and connectors, and hot glue.<br />
[[File:Aspirator.PNG|center|500px]]<br />
<br />
You can easily build a GRIN lens holder by using a suction system to hold the lens in place as it is dropped into the brain for implantation. To build, start with two 1ml micropipette tips. Cut the first so that the tip is just larger than the diameter of the GRIN lens you will be using. Cut the second tip so that the end is just smaller than the size of the GRIN lens. Connect this end to a 1ml syringe and connect to a vacuum line. This will then hold the lens in place during implantation. You can then use tape to attach this holder to a stereotax. Another solution we have used is a basic drill chuck, but this is only good for large (2mm) lenses.<br />
[[File:LensHolder.PNG|center|500px]]<br />
<br />
=== Skull Preparation ===<br />
You must first prepare the skull for implantation. We recommend shaving the head area and sterilizing with 3 alternating betadine and ethanol scrubs. Next, remove the scalp with scissors and clean the skull with hydrogen peroxide and saline. Scrape and score the skull to increase the bond with the glue. We also detach the neck muscle from the skull in order to reduce pull on the skull and prevent muscle growth that can make the implant less stable. Finally we insert a skull screw on the opposite side of the skull to enhance the stability of the implant.<br />
<br />
=== Craniotomy and Aspiration ===<br />
Using a stereotax, align the skull and find the location for the implant. Using a drill, make 4 guide holes to outline the lens placement. For CA1, we recommend offsetting the lens 0.5mm to the medial side of the virus injection to prevent imaging any damage induced by the virus injection. Connect the guide holes to create a circular craniotomy and peel off the circular skull fragment. Do not worry about blood as the top layers of cortex will be removed by aspiration. Make absolutely sure that the craniotomy is at least the size of your GRIN lens! Cover with cortex buffer and scrape around the outside of the craniotomy to remove any excess bone or dura. <br />
<br />
While keeping a constant flow of cortex buffer use the aspirator to slowly remove the cortex above the hippocampus. Continue removing cortex until you reach the white striations of the corpus callosum. Very slowly and carefully remove the horizontal white striations of the corpus callosum until you reach the striations that go vertical. When you reach these striations, stop aspirating and clean up the sides so that the lens can be placed into the brain. Continue washing with cortex buffer until all bleeding stops and keep the brain constantly flushed with cortex buffer.<br />
<br />
=== Lens implantation ===<br />
Attach GRIN lens to lens holder on stereotax. Align the lens with the top of the skull at the most posterior part of the craniotomy and quickly insert the lens 1.35 mm below the top of the skull. Remove any excess liquid with the aspirator. Connect the outside of the GRIN lens with the skull and skull screw using cyanoacrylate glue and let fully dry or use a glue accelerator. Remove the lens holder by simply removing the vacuum suction (you can just pinch the tubing) and withdrawing the holder. Next, cover the entire skull with cyanoacrylate glue, and then cover with dental cement. Let dry completely and cover with Kwik-Sil to protect the lens. Remove from stereotax and allow the animal to recover. Give the animal amoxicillin (or equivalent) for 7 days, and daily injections of carprofen and dexamethasone for 7 days.<br />
<br />
=== Important tips to remember ===<br />
*Try to keep the surgery as sterile as possible. Infections will destroy imaging and the ability to keep the same cells over many days.<br />
<br />
*Try to keep isoflurane as low as possible. Surgeries can take 2-6 hours depending on experience.<br />
<br />
*Aspirations will take a lot of practice. Go slow at first.<br />
<br />
*Depth will be trial and error. If the first few animals don't give good images, check the depth using histology and adjust accordingly.<br />
<br />
<br />
Please feel free to add any tips that you have!<br />
<br />
== Baseplating Protocol ==<br />
<br />
<br />
=== Basic Equipment needed ===<br />
You will only need basic surgical equipment to baseplate. This includes a mouse stereotax, an isoflurane vaporizer, surgical heat pad, stereo surgical microscope, light source, and a dental drill. <br />
<br />
You will also need cyanoacrylate glue, fine forceps, double distilled water, lens paper, and dental cement<br />
=== Baseplate Preparation ===<br />
Before affixing the baseplate on an animal, you will need to prepare the baseplate. <br />
<br />
The first step is to insert magnets into the appropriate holes on the baseplate. To ensure that the polarity of the magnets is correct throughout the baseplate preparation, use a reference Miniscope to hold onto the magnets. Mark the top magnet with a sharpie, as seen in the picture below. Next, put the top magnet on the head of a screwdriver with the marked side facing away from the screwdriver. Apply cyanoacrylate glue on the edge of the magnet, then push the magnet through the appropriate hole. Ensure that the magnet goes through the hole completely and the top side of the baseplate is flat, as the quality of imaging relies on the Miniscope sitting flush on the baseplate. Repeat this procedure for the next 2 magnets. Finally, put a coat of cyanoacrylate glue on the bottom side of the baseplate and leave the baseplate to dry.<br />
<br />
Once the baseplate has dried, score the bottom and the sides of the baseplate using a dental drill to allow the dental glue to adhere the baseplate and skull well.<br />
<br />
=== Cap Preparation ===<br />
Similar to the baseplate preparation, use a reference Miniscope to hold onto the magnets. Mark the top magnet with a sharpie, and put the magnet on the head of a screwdriver with the marked side facing toward the screwdriver. Coat the magnet with cyanoacrylate glue and push the magnet into the appropriate hole. Repeat for the remaining two magnets.<br />
=== Baseplating ===<br />
Connect a Miniscope to a DAQ and initialize the Miniscope software on a computer. Attach your prepared baseplate onto the bottom of the Miniscope using a set screw.<br />
<br />
Using a stereotax and isoflurane vaporizer, anesthetize the animal and fix ear bars until the animal is stable. Carefully remove the Kwik-Sil off the top of the animal’s GRIN lens with blunt forceps being sure not to scratch the lens. Under a stereo surgical microscope, check to see if there is debris on the top of the GRIN lens. If there is, dampen a sheet of lens paper using double distilled water. Wipe the top of the GRIN lens using fine forceps and lens paper, being sure to only wipe laterally and not to apply direct downward pressure.<br />
<br />
After the top of the GRIN lens is clean, explore the field of view with the Miniscope. Adjust the LED brightness and focus accordingly. We advise that you use maximum gain and low LED excitation as to not photobleach the brain during baseplating. After finding your ideal view, practice taking the Miniscope off and on and finding the same view. <br />
<br />
When you feel comfortable repeatedly finding the view with the Miniscope, apply dental cement in a “C” pattern around the GRIN lens, making sure not to touch the lens itself. It will be easier if your first application of dental cement is more viscous. Put the Miniscope on top of the lens and find your ideal view. Hold the Miniscope steadily as the first application of dental cement dries, making sure that your view does not change. <br />
After the first application of dental cement dries, mix more dental cement and apply it on the posterior edge of the baseplate, making sure it seals to the skull. At this point, you can also build up dental cement along the wall of the baseplate on the animal’s right side. Be sure to avoid applying dental cement onto the camera itself and adhering it to the baseplate. Once the dental cement dries and you feel confident that the baseplate will no longer shift, carefully remove the Miniscope from the baseplate by unscrewing the set screw and lifting it off. <br />
<br />
Apply dental cement to the anterior edge of the baseplate and fill in the gaps between the baseplate and skull on the animal’s left side. Allow the dental cement to dry and attach the plastic cap onto the top of the baseplate with a set screw.</div>Chrisl1026http://miniscope.org/index.php?title=Surgery_Protocol&diff=1509Surgery Protocol2016-09-21T02:14:24Z<p>Chrisl1026: /* Baseplate Preparation */</p>
<hr />
<div>The following is a basic outline of our procedure for implanting a GRIN lens above the hippocampus for CA1 imaging. For videos of surgery and baseplating, see the [[Online Workshop]]. Also, see the recent Nature Protocols paper from the Stuber Lab ([http://www.nature.com/nprot/journal/v11/n3/full/nprot.2016.021.html link]) that has a very detailed description of the surgery. <br />
<br />
In order to image, animals will need to undergo three surgical procedures: a virus injection, a GRIN lens implantation, and baseplate attachment.<br />
<br />
== Virus Injection ==<br />
Before implanting the GRIN lens, you will need to inject a fluorescent indicator such as GCaMP6. We have generally used AAV1.Syn.GCaMP6f.WPRE.SV40 from Penn Vector (Item number AV-1-PV2822) and had great success in dorsal CA1. You may also be able to use GCaMP6 transgenic mice, but we have not tested whether any of the newer lines are bright enough to be imaged.<br />
<br />
== GRIN Lens Implantation==<br />
<br />
=== Basic Equipment Needed ===<br />
You will only need basic surgical equipment to perform GRIN lens implantation surgeries. This includes a mouse stereotax, an isoflurane vaporizor, surgical heat pad, stereo surgical microscope, light source, dental drill, and bead sterilizer. For recommended equipment, see the master [https://docs.google.com/spreadsheets/d/12H71DU2QX8d7efUE4yNuikBEiIzKaXjYqdc0A-oLNSw/edit?usp=sharing parts list].<br />
<br />
You will need the following basic tools and supplies: fine forceps, blunt forceps, fine scissors, scalpel and blade, small skull screws, drill bits, cyanoacrylate glue, dental cement, Kwik-Sil, and [[cortex buffer]]<br />
<br />
=== Aspirator and Lens Holder ===<br />
You will need an aspirator to remove cortex above the hippocampus. This can be built very simply with a vacuum line (or pump), a liquid trap, 1 ml syringe (with a hole to control suction), blunt needles, tubing and connectors, and hot glue.<br />
[[File:Aspirator.PNG|center|500px]]<br />
<br />
You can easily build a GRIN lens holder by using a suction system to hold the lens in place as it is dropped into the brain for implantation. To build, start with two 1ml micropipette tips. Cut the first so that the tip is just larger than the diameter of the GRIN lens you will be using. Cut the second tip so that the end is just smaller than the size of the GRIN lens. Connect this end to a 1ml syringe and connect to a vacuum line. This will then hold the lens in place during implantation. You can then use tape to attach this holder to a stereotax. Another solution we have used is a basic drill chuck, but this is only good for large (2mm) lenses.<br />
[[File:LensHolder.PNG|center|500px]]<br />
<br />
=== Skull Preparation ===<br />
You must first prepare the skull for implantation. We recommend shaving the head area and sterilizing with 3 alternating betadine and ethanol scrubs. Next, remove the scalp with scissors and clean the skull with hydrogen peroxide and saline. Scrape and score the skull to increase the bond with the glue. We also detach the neck muscle from the skull in order to reduce pull on the skull and prevent muscle growth that can make the implant less stable. Finally we insert a skull screw on the opposite side of the skull to enhance the stability of the implant.<br />
<br />
=== Craniotomy and Aspiration ===<br />
Using a stereotax, align the skull and find the location for the implant. Using a drill, make 4 guide holes to outline the lens placement. For CA1, we recommend offsetting the lens 0.5mm to the medial side of the virus injection to prevent imaging any damage induced by the virus injection. Connect the guide holes to create a circular craniotomy and peel off the circular skull fragment. Do not worry about blood as the top layers of cortex will be removed by aspiration. Make absolutely sure that the craniotomy is at least the size of your GRIN lens! Cover with cortex buffer and scrape around the outside of the craniotomy to remove any excess bone or dura. <br />
<br />
While keeping a constant flow of cortex buffer use the aspirator to slowly remove the cortex above the hippocampus. Continue removing cortex until you reach the white striations of the corpus callosum. Very slowly and carefully remove the horizontal white striations of the corpus callosum until you reach the striations that go vertical. When you reach these striations, stop aspirating and clean up the sides so that the lens can be placed into the brain. Continue washing with cortex buffer until all bleeding stops and keep the brain constantly flushed with cortex buffer.<br />
<br />
=== Lens implantation ===<br />
Attach GRIN lens to lens holder on stereotax. Align the lens with the top of the skull at the most posterior part of the craniotomy and quickly insert the lens 1.35 mm below the top of the skull. Remove any excess liquid with the aspirator. Connect the outside of the GRIN lens with the skull and skull screw using cyanoacrylate glue and let fully dry or use a glue accelerator. Remove the lens holder by simply removing the vacuum suction (you can just pinch the tubing) and withdrawing the holder. Next, cover the entire skull with cyanoacrylate glue, and then cover with dental cement. Let dry completely and cover with Kwik-Sil to protect the lens. Remove from stereotax and allow the animal to recover. Give the animal amoxicillin (or equivalent) for 7 days, and daily injections of carprofen and dexamethasone for 7 days.<br />
<br />
=== Important tips to remember ===<br />
*Try to keep the surgery as sterile as possible. Infections will destroy imaging and the ability to keep the same cells over many days.<br />
<br />
*Try to keep isoflurane as low as possible. Surgeries can take 2-6 hours depending on experience.<br />
<br />
*Aspirations will take a lot of practice. Go slow at first.<br />
<br />
*Depth will be trial and error. If the first few animals don't give good images, check the depth using histology and adjust accordingly.<br />
<br />
<br />
Please feel free to add any tips that you have!<br />
<br />
== Baseplating Protocol ==<br />
<br />
<br />
=== Basic Equipment needed ===<br />
You will only need basic surgical equipment to baseplate. This includes a mouse stereotax, an isoflurane vaporizer, surgical heat pad, stereo surgical microscope, light source, and a dental drill. <br />
<br />
You will also need cyanoacrylate glue, fine forceps, double distilled water, lens paper, and dental cement<br />
=== Baseplate Preparation ===<br />
Before affixing the baseplate on an animal, you will need to prepare the baseplate. <br />
<br />
The first step is to insert magnets into the appropriate holes on the baseplate. To ensure that the polarity of the magnets is correct throughout the baseplate preparation, use a reference Miniscope to hold onto the magnets. Mark the top magnet with a sharpie, as seen in the picture below. Next, put the top magnet on the head of a screwdriver with the marked side facing away from the screwdriver. Apply cyanoacrylate glue on the edge of the magnet, then push the magnet through the appropriate hole. Ensure that the magnet goes through the hole completely and the top side of the baseplate is flat, as the quality of imaging relies on the Miniscope sitting flush on the baseplate. Repeat this procedure for the next 2 magnets. Finally, put a coat of cyanoacrylate glue on the bottom side of the baseplate and leave the baseplate to dry.<br />
<br />
Once the baseplate has dried, score the bottom and the sides of the baseplate using a dental drill to allow the dental glue to adhere the baseplate and skull well.<br />
<br />
[[File:BP_Prep_1.jpeg]] [[File:BP_Prep_2.jpeg]] [[File:BP_Prep_3.jpeg]] [[File:BP_Prep_4.jpeg]] [[File:BP_Prep_5.jpeg]]<br />
<br />
=== Cap Preparation ===<br />
Similar to the baseplate preparation, use a reference Miniscope to hold onto the magnets. Mark the top magnet with a sharpie, and put the magnet on the head of a screwdriver with the marked side facing toward the screwdriver. Coat the magnet with cyanoacrylate glue and push the magnet into the appropriate hole. Repeat for the remaining two magnets.<br />
=== Baseplating ===<br />
Connect a Miniscope to a DAQ and initialize the Miniscope software on a computer. Attach your prepared baseplate onto the bottom of the Miniscope using a set screw.<br />
<br />
Using a stereotax and isoflurane vaporizer, anesthetize the animal and fix ear bars until the animal is stable. Carefully remove the Kwik-Sil off the top of the animal’s GRIN lens with blunt forceps being sure not to scratch the lens. Under a stereo surgical microscope, check to see if there is debris on the top of the GRIN lens. If there is, dampen a sheet of lens paper using double distilled water. Wipe the top of the GRIN lens using fine forceps and lens paper, being sure to only wipe laterally and not to apply direct downward pressure.<br />
<br />
After the top of the GRIN lens is clean, explore the field of view with the Miniscope. Adjust the LED brightness and focus accordingly. We advise that you use maximum gain and low LED excitation as to not photobleach the brain during baseplating. After finding your ideal view, practice taking the Miniscope off and on and finding the same view. <br />
<br />
When you feel comfortable repeatedly finding the view with the Miniscope, apply dental cement in a “C” pattern around the GRIN lens, making sure not to touch the lens itself. It will be easier if your first application of dental cement is more viscous. Put the Miniscope on top of the lens and find your ideal view. Hold the Miniscope steadily as the first application of dental cement dries, making sure that your view does not change. <br />
After the first application of dental cement dries, mix more dental cement and apply it on the posterior edge of the baseplate, making sure it seals to the skull. At this point, you can also build up dental cement along the wall of the baseplate on the animal’s right side. Be sure to avoid applying dental cement onto the camera itself and adhering it to the baseplate. Once the dental cement dries and you feel confident that the baseplate will no longer shift, carefully remove the Miniscope from the baseplate by unscrewing the set screw and lifting it off. <br />
<br />
Apply dental cement to the anterior edge of the baseplate and fill in the gaps between the baseplate and skull on the animal’s left side. Allow the dental cement to dry and attach the plastic cap onto the top of the baseplate with a set screw.</div>Chrisl1026http://miniscope.org/index.php?title=Surgery_Protocol&diff=1508Surgery Protocol2016-09-20T23:43:09Z<p>Chrisl1026: /* Baseplating Protocol */</p>
<hr />
<div>The following is a basic outline of our procedure for implanting a GRIN lens above the hippocampus for CA1 imaging. For videos of surgery and baseplating, see the [[Online Workshop]]. Also, see the recent Nature Protocols paper from the Stuber Lab ([http://www.nature.com/nprot/journal/v11/n3/full/nprot.2016.021.html link]) that has a very detailed description of the surgery. <br />
<br />
In order to image, animals will need to undergo three surgical procedures: a virus injection, a GRIN lens implantation, and baseplate attachment.<br />
<br />
== Virus Injection ==<br />
Before implanting the GRIN lens, you will need to inject a fluorescent indicator such as GCaMP6. We have generally used AAV1.Syn.GCaMP6f.WPRE.SV40 from Penn Vector (Item number AV-1-PV2822) and had great success in dorsal CA1. You may also be able to use GCaMP6 transgenic mice, but we have not tested whether any of the newer lines are bright enough to be imaged.<br />
<br />
== GRIN Lens Implantation==<br />
<br />
=== Basic Equipment Needed ===<br />
You will only need basic surgical equipment to perform GRIN lens implantation surgeries. This includes a mouse stereotax, an isoflurane vaporizor, surgical heat pad, stereo surgical microscope, light source, dental drill, and bead sterilizer. For recommended equipment, see the master [https://docs.google.com/spreadsheets/d/12H71DU2QX8d7efUE4yNuikBEiIzKaXjYqdc0A-oLNSw/edit?usp=sharing parts list].<br />
<br />
You will need the following basic tools and supplies: fine forceps, blunt forceps, fine scissors, scalpel and blade, small skull screws, drill bits, cyanoacrylate glue, dental cement, Kwik-Sil, and [[cortex buffer]]<br />
<br />
=== Aspirator and Lens Holder ===<br />
You will need an aspirator to remove cortex above the hippocampus. This can be built very simply with a vacuum line (or pump), a liquid trap, 1 ml syringe (with a hole to control suction), blunt needles, tubing and connectors, and hot glue.<br />
[[File:Aspirator.PNG|center|500px]]<br />
<br />
You can easily build a GRIN lens holder by using a suction system to hold the lens in place as it is dropped into the brain for implantation. To build, start with two 1ml micropipette tips. Cut the first so that the tip is just larger than the diameter of the GRIN lens you will be using. Cut the second tip so that the end is just smaller than the size of the GRIN lens. Connect this end to a 1ml syringe and connect to a vacuum line. This will then hold the lens in place during implantation. You can then use tape to attach this holder to a stereotax. Another solution we have used is a basic drill chuck, but this is only good for large (2mm) lenses.<br />
[[File:LensHolder.PNG|center|500px]]<br />
<br />
=== Skull Preparation ===<br />
You must first prepare the skull for implantation. We recommend shaving the head area and sterilizing with 3 alternating betadine and ethanol scrubs. Next, remove the scalp with scissors and clean the skull with hydrogen peroxide and saline. Scrape and score the skull to increase the bond with the glue. We also detach the neck muscle from the skull in order to reduce pull on the skull and prevent muscle growth that can make the implant less stable. Finally we insert a skull screw on the opposite side of the skull to enhance the stability of the implant.<br />
<br />
=== Craniotomy and Aspiration ===<br />
Using a stereotax, align the skull and find the location for the implant. Using a drill, make 4 guide holes to outline the lens placement. For CA1, we recommend offsetting the lens 0.5mm to the medial side of the virus injection to prevent imaging any damage induced by the virus injection. Connect the guide holes to create a circular craniotomy and peel off the circular skull fragment. Do not worry about blood as the top layers of cortex will be removed by aspiration. Make absolutely sure that the craniotomy is at least the size of your GRIN lens! Cover with cortex buffer and scrape around the outside of the craniotomy to remove any excess bone or dura. <br />
<br />
While keeping a constant flow of cortex buffer use the aspirator to slowly remove the cortex above the hippocampus. Continue removing cortex until you reach the white striations of the corpus callosum. Very slowly and carefully remove the horizontal white striations of the corpus callosum until you reach the striations that go vertical. When you reach these striations, stop aspirating and clean up the sides so that the lens can be placed into the brain. Continue washing with cortex buffer until all bleeding stops and keep the brain constantly flushed with cortex buffer.<br />
<br />
=== Lens implantation ===<br />
Attach GRIN lens to lens holder on stereotax. Align the lens with the top of the skull at the most posterior part of the craniotomy and quickly insert the lens 1.35 mm below the top of the skull. Remove any excess liquid with the aspirator. Connect the outside of the GRIN lens with the skull and skull screw using cyanoacrylate glue and let fully dry or use a glue accelerator. Remove the lens holder by simply removing the vacuum suction (you can just pinch the tubing) and withdrawing the holder. Next, cover the entire skull with cyanoacrylate glue, and then cover with dental cement. Let dry completely and cover with Kwik-Sil to protect the lens. Remove from stereotax and allow the animal to recover. Give the animal amoxicillin (or equivalent) for 7 days, and daily injections of carprofen and dexamethasone for 7 days.<br />
<br />
=== Important tips to remember ===<br />
*Try to keep the surgery as sterile as possible. Infections will destroy imaging and the ability to keep the same cells over many days.<br />
<br />
*Try to keep isoflurane as low as possible. Surgeries can take 2-6 hours depending on experience.<br />
<br />
*Aspirations will take a lot of practice. Go slow at first.<br />
<br />
*Depth will be trial and error. If the first few animals don't give good images, check the depth using histology and adjust accordingly.<br />
<br />
<br />
Please feel free to add any tips that you have!<br />
<br />
== Baseplating Protocol ==<br />
<br />
<br />
=== Basic Equipment needed ===<br />
You will only need basic surgical equipment to baseplate. This includes a mouse stereotax, an isoflurane vaporizer, surgical heat pad, stereo surgical microscope, light source, and a dental drill. <br />
<br />
You will also need cyanoacrylate glue, fine forceps, double distilled water, lens paper, and dental cement<br />
=== Baseplate Preparation ===<br />
Before affixing the baseplate on an animal, you will need to prepare the baseplate. <br />
<br />
The first step is to insert magnets into the appropriate holes on the baseplate. To ensure that the polarity of the magnets is correct throughout the baseplate preparation, use a reference Miniscope to hold onto the magnets. Mark the top magnet with a sharpie, as seen in the picture below. Next, put the top magnet on the head of a screwdriver with the marked side facing away from the screwdriver. Apply cyanoacrylate glue on the edge of the magnet, then push the magnet through the appropriate hole. Ensure that the magnet goes through the hole completely and the top side of the baseplate is flat, as the quality of imaging relies on the Miniscope sitting flush on the baseplate. Repeat this procedure for the next 2 magnets. Finally, put a coat of cyanoacrylate glue on the bottom side of the baseplate and leave the baseplate to dry.<br />
<br />
Once the baseplate has dried, score the bottom and the sides of the baseplate using a dental drill to allow the dental glue to adhere the baseplate and skull well. <br />
=== Cap Preparation ===<br />
Similar to the baseplate preparation, use a reference Miniscope to hold onto the magnets. Mark the top magnet with a sharpie, and put the magnet on the head of a screwdriver with the marked side facing toward the screwdriver. Coat the magnet with cyanoacrylate glue and push the magnet into the appropriate hole. Repeat for the remaining two magnets.<br />
=== Baseplating ===<br />
Connect a Miniscope to a DAQ and initialize the Miniscope software on a computer. Attach your prepared baseplate onto the bottom of the Miniscope using a set screw.<br />
<br />
Using a stereotax and isoflurane vaporizer, anesthetize the animal and fix ear bars until the animal is stable. Carefully remove the Kwik-Sil off the top of the animal’s GRIN lens with blunt forceps being sure not to scratch the lens. Under a stereo surgical microscope, check to see if there is debris on the top of the GRIN lens. If there is, dampen a sheet of lens paper using double distilled water. Wipe the top of the GRIN lens using fine forceps and lens paper, being sure to only wipe laterally and not to apply direct downward pressure.<br />
<br />
After the top of the GRIN lens is clean, explore the field of view with the Miniscope. Adjust the LED brightness and focus accordingly. We advise that you use maximum gain and low LED excitation as to not photobleach the brain during baseplating. After finding your ideal view, practice taking the Miniscope off and on and finding the same view. <br />
<br />
When you feel comfortable repeatedly finding the view with the Miniscope, apply dental cement in a “C” pattern around the GRIN lens, making sure not to touch the lens itself. It will be easier if your first application of dental cement is more viscous. Put the Miniscope on top of the lens and find your ideal view. Hold the Miniscope steadily as the first application of dental cement dries, making sure that your view does not change. <br />
After the first application of dental cement dries, mix more dental cement and apply it on the posterior edge of the baseplate, making sure it seals to the skull. At this point, you can also build up dental cement along the wall of the baseplate on the animal’s right side. Be sure to avoid applying dental cement onto the camera itself and adhering it to the baseplate. Once the dental cement dries and you feel confident that the baseplate will no longer shift, carefully remove the Miniscope from the baseplate by unscrewing the set screw and lifting it off. <br />
<br />
Apply dental cement to the anterior edge of the baseplate and fill in the gaps between the baseplate and skull on the animal’s left side. Allow the dental cement to dry and attach the plastic cap onto the top of the baseplate with a set screw.</div>Chrisl1026http://miniscope.org/index.php?title=Surgery_Protocol&diff=1507Surgery Protocol2016-09-20T21:08:13Z<p>Chrisl1026: /* Baseplating Protocol */</p>
<hr />
<div>The following is a basic outline of our procedure for implanting a GRIN lens above the hippocampus for CA1 imaging. For videos of surgery and baseplating, see the [[Online Workshop]]. Also, see the recent Nature Protocols paper from the Stuber Lab ([http://www.nature.com/nprot/journal/v11/n3/full/nprot.2016.021.html link]) that has a very detailed description of the surgery. <br />
<br />
In order to image, animals will need to undergo three surgical procedures: a virus injection, a GRIN lens implantation, and baseplate attachment.<br />
<br />
== Virus Injection ==<br />
Before implanting the GRIN lens, you will need to inject a fluorescent indicator such as GCaMP6. We have generally used AAV1.Syn.GCaMP6f.WPRE.SV40 from Penn Vector (Item number AV-1-PV2822) and had great success in dorsal CA1. You may also be able to use GCaMP6 transgenic mice, but we have not tested whether any of the newer lines are bright enough to be imaged.<br />
<br />
== GRIN Lens Implantation==<br />
<br />
=== Basic Equipment Needed ===<br />
You will only need basic surgical equipment to perform GRIN lens implantation surgeries. This includes a mouse stereotax, an isoflurane vaporizor, surgical heat pad, stereo surgical microscope, light source, dental drill, and bead sterilizer. For recommended equipment, see the master [https://docs.google.com/spreadsheets/d/12H71DU2QX8d7efUE4yNuikBEiIzKaXjYqdc0A-oLNSw/edit?usp=sharing parts list].<br />
<br />
You will need the following basic tools and supplies: fine forceps, blunt forceps, fine scissors, scalpel and blade, small skull screws, drill bits, cyanoacrylate glue, dental cement, Kwik-Sil, and [[cortex buffer]]<br />
<br />
=== Aspirator and Lens Holder ===<br />
You will need an aspirator to remove cortex above the hippocampus. This can be built very simply with a vacuum line (or pump), a liquid trap, 1 ml syringe (with a hole to control suction), blunt needles, tubing and connectors, and hot glue.<br />
[[File:Aspirator.PNG|center|500px]]<br />
<br />
You can easily build a GRIN lens holder by using a suction system to hold the lens in place as it is dropped into the brain for implantation. To build, start with two 1ml micropipette tips. Cut the first so that the tip is just larger than the diameter of the GRIN lens you will be using. Cut the second tip so that the end is just smaller than the size of the GRIN lens. Connect this end to a 1ml syringe and connect to a vacuum line. This will then hold the lens in place during implantation. You can then use tape to attach this holder to a stereotax. Another solution we have used is a basic drill chuck, but this is only good for large (2mm) lenses.<br />
[[File:LensHolder.PNG|center|500px]]<br />
<br />
=== Skull Preparation ===<br />
You must first prepare the skull for implantation. We recommend shaving the head area and sterilizing with 3 alternating betadine and ethanol scrubs. Next, remove the scalp with scissors and clean the skull with hydrogen peroxide and saline. Scrape and score the skull to increase the bond with the glue. We also detach the neck muscle from the skull in order to reduce pull on the skull and prevent muscle growth that can make the implant less stable. Finally we insert a skull screw on the opposite side of the skull to enhance the stability of the implant.<br />
<br />
=== Craniotomy and Aspiration ===<br />
Using a stereotax, align the skull and find the location for the implant. Using a drill, make 4 guide holes to outline the lens placement. For CA1, we recommend offsetting the lens 0.5mm to the medial side of the virus injection to prevent imaging any damage induced by the virus injection. Connect the guide holes to create a circular craniotomy and peel off the circular skull fragment. Do not worry about blood as the top layers of cortex will be removed by aspiration. Make absolutely sure that the craniotomy is at least the size of your GRIN lens! Cover with cortex buffer and scrape around the outside of the craniotomy to remove any excess bone or dura. <br />
<br />
While keeping a constant flow of cortex buffer use the aspirator to slowly remove the cortex above the hippocampus. Continue removing cortex until you reach the white striations of the corpus callosum. Very slowly and carefully remove the horizontal white striations of the corpus callosum until you reach the striations that go vertical. When you reach these striations, stop aspirating and clean up the sides so that the lens can be placed into the brain. Continue washing with cortex buffer until all bleeding stops and keep the brain constantly flushed with cortex buffer.<br />
<br />
=== Lens implantation ===<br />
Attach GRIN lens to lens holder on stereotax. Align the lens with the top of the skull at the most posterior part of the craniotomy and quickly insert the lens 1.35 mm below the top of the skull. Remove any excess liquid with the aspirator. Connect the outside of the GRIN lens with the skull and skull screw using cyanoacrylate glue and let fully dry or use a glue accelerator. Remove the lens holder by simply removing the vacuum suction (you can just pinch the tubing) and withdrawing the holder. Next, cover the entire skull with cyanoacrylate glue, and then cover with dental cement. Let dry completely and cover with Kwik-Sil to protect the lens. Remove from stereotax and allow the animal to recover. Give the animal amoxicillin (or equivalent) for 7 days, and daily injections of carprofen and dexamethasone for 7 days.<br />
<br />
=== Important tips to remember ===<br />
*Try to keep the surgery as sterile as possible. Infections will destroy imaging and the ability to keep the same cells over many days.<br />
<br />
*Try to keep isoflurane as low as possible. Surgeries can take 2-6 hours depending on experience.<br />
<br />
*Aspirations will take a lot of practice. Go slow at first.<br />
<br />
*Depth will be trial and error. If the first few animals don't give good images, check the depth using histology and adjust accordingly.<br />
<br />
<br />
Please feel free to add any tips that you have!<br />
<br />
== Baseplating Protocol ==<br />
<br />
<br />
=== Basic Equipment needed ===<br />
<br />
You will only need basic surgical equipment to baseplate. This includes a mouse stereotax, an isoflurane vaporizer, surgical heat pad, stereo surgical microscope, light source, and a dental drill. <br />
<br />
You will also need cyanoacrylate glue, fine forceps, double distilled water, lens paper, and dental cement<br />
<br />
<br />
=== Baseplate Preparation ===<br />
<br />
Before affixing the baseplate on an animal, you will need to prepare the baseplate. <br />
<br />
The first step is to insert magnets into the appropriate holes on the baseplate. To ensure that the polarity of the magnets is correct throughout the baseplate preparation, use a reference Miniscope to hold onto the magnets. Mark the top magnet with a sharpie, as seen in the picture below. Next, put the top magnet on the head of a screwdriver with the marked side facing away from the screwdriver. Apply cyanoacrylate glue on the edge of the magnet, then push the magnet through the appropriate hole. Ensure that the magnet goes through the hole completely and the top side of the baseplate is flat, as the quality of imaging relies on the Miniscope sitting flush on the baseplate. Repeat this procedure for the next 2 magnets. Finally, put a coat of cyanoacrylate glue on the bottom side of the baseplate and leave the baseplate to dry.<br />
<br />
Once the baseplate has dried, score the bottom and the sides of the baseplate using a dental drill to allow the dental glue to adhere the baseplate and skull well. <br />
<br />
<br />
<br />
=== Cap Preparation ===<br />
<br />
Similar to the baseplate preparation, use a reference Miniscope to hold onto the magnets. Mark the top magnet with a sharpie, and put the magnet on the head of a screwdriver with the marked side facing toward the screwdriver. Coat the magnet with cyanoacrylate glue and push the magnet into the appropriate hole. Repeat for the remaining two magnets.<br />
<br />
<br />
=== Baseplating ===<br />
<br />
Connect a Miniscope to a DAQ and initialize the Miniscope software on a computer. Attach your prepared baseplate onto the bottom of the Miniscope using a set screw.<br />
<br />
Using a stereotax and isoflurane vaporizer, anesthetize the animal and fix ear bars until the animal is stable. Carefully remove the Kwik-Sil off the top of the animal’s GRIN lens with blunt forceps being sure not to scratch the lens. Under a stereo surgical microscope, check to see if there is debris on the top of the GRIN lens. If there is, dampen a sheet of lens paper using double distilled water. Wipe the top of the GRIN lens using fine forceps and lens paper, being sure to only wipe laterally and not to apply direct downward pressure.<br />
<br />
After the top of the GRIN lens is clean, explore the field of view with the Miniscope. Adjust the LED brightness and focus accordingly. We advise that you use maximum gain and low LED excitation as to not photobleach the brain during baseplating. After finding your ideal view, practice taking the Miniscope off and on and finding the same view. <br />
<br />
When you feel comfortable repeatedly finding the view with the Miniscope, apply dental cement in a “C” pattern around the GRIN lens, making sure not to touch the lens itself. It will be easier if your first application of dental cement is more viscous. Put the Miniscope on top of the lens and find your ideal view. Hold the Miniscope steadily as the first application of dental cement dries, making sure that your view does not change. <br />
After the first application of dental cement dries, mix more dental cement and apply it on the posterior edge of the baseplate, making sure it seals to the skull. At this point, you can also build up dental cement along the wall of the baseplate on the animal’s right side. Be sure to avoid applying dental cement onto the camera itself and adhering it to the baseplate. Once the dental cement dries and you feel confident that the baseplate will no longer shift, carefully remove the Miniscope from the baseplate by unscrewing the set screw and lifting it off. <br />
<br />
Apply dental cement to the anterior edge of the baseplate and fill in the gaps between the baseplate and skull on the animal’s left side. Allow the dental cement to dry and attach the plastic cap onto the top of the baseplate with a set screw.</div>Chrisl1026http://miniscope.org/index.php?title=Surgery_Protocol&diff=1506Surgery Protocol2016-09-20T21:07:15Z<p>Chrisl1026: /* Baseplating Protocol */</p>
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<div>The following is a basic outline of our procedure for implanting a GRIN lens above the hippocampus for CA1 imaging. For videos of surgery and baseplating, see the [[Online Workshop]]. Also, see the recent Nature Protocols paper from the Stuber Lab ([http://www.nature.com/nprot/journal/v11/n3/full/nprot.2016.021.html link]) that has a very detailed description of the surgery. <br />
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In order to image, animals will need to undergo three surgical procedures: a virus injection, a GRIN lens implantation, and baseplate attachment.<br />
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== Virus Injection ==<br />
Before implanting the GRIN lens, you will need to inject a fluorescent indicator such as GCaMP6. We have generally used AAV1.Syn.GCaMP6f.WPRE.SV40 from Penn Vector (Item number AV-1-PV2822) and had great success in dorsal CA1. You may also be able to use GCaMP6 transgenic mice, but we have not tested whether any of the newer lines are bright enough to be imaged.<br />
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== GRIN Lens Implantation==<br />
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=== Basic Equipment Needed ===<br />
You will only need basic surgical equipment to perform GRIN lens implantation surgeries. This includes a mouse stereotax, an isoflurane vaporizor, surgical heat pad, stereo surgical microscope, light source, dental drill, and bead sterilizer. For recommended equipment, see the master [https://docs.google.com/spreadsheets/d/12H71DU2QX8d7efUE4yNuikBEiIzKaXjYqdc0A-oLNSw/edit?usp=sharing parts list].<br />
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You will need the following basic tools and supplies: fine forceps, blunt forceps, fine scissors, scalpel and blade, small skull screws, drill bits, cyanoacrylate glue, dental cement, Kwik-Sil, and [[cortex buffer]]<br />
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=== Aspirator and Lens Holder ===<br />
You will need an aspirator to remove cortex above the hippocampus. This can be built very simply with a vacuum line (or pump), a liquid trap, 1 ml syringe (with a hole to control suction), blunt needles, tubing and connectors, and hot glue.<br />
[[File:Aspirator.PNG|center|500px]]<br />
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You can easily build a GRIN lens holder by using a suction system to hold the lens in place as it is dropped into the brain for implantation. To build, start with two 1ml micropipette tips. Cut the first so that the tip is just larger than the diameter of the GRIN lens you will be using. Cut the second tip so that the end is just smaller than the size of the GRIN lens. Connect this end to a 1ml syringe and connect to a vacuum line. This will then hold the lens in place during implantation. You can then use tape to attach this holder to a stereotax. Another solution we have used is a basic drill chuck, but this is only good for large (2mm) lenses.<br />
[[File:LensHolder.PNG|center|500px]]<br />
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=== Skull Preparation ===<br />
You must first prepare the skull for implantation. We recommend shaving the head area and sterilizing with 3 alternating betadine and ethanol scrubs. Next, remove the scalp with scissors and clean the skull with hydrogen peroxide and saline. Scrape and score the skull to increase the bond with the glue. We also detach the neck muscle from the skull in order to reduce pull on the skull and prevent muscle growth that can make the implant less stable. Finally we insert a skull screw on the opposite side of the skull to enhance the stability of the implant.<br />
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=== Craniotomy and Aspiration ===<br />
Using a stereotax, align the skull and find the location for the implant. Using a drill, make 4 guide holes to outline the lens placement. For CA1, we recommend offsetting the lens 0.5mm to the medial side of the virus injection to prevent imaging any damage induced by the virus injection. Connect the guide holes to create a circular craniotomy and peel off the circular skull fragment. Do not worry about blood as the top layers of cortex will be removed by aspiration. Make absolutely sure that the craniotomy is at least the size of your GRIN lens! Cover with cortex buffer and scrape around the outside of the craniotomy to remove any excess bone or dura. <br />
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While keeping a constant flow of cortex buffer use the aspirator to slowly remove the cortex above the hippocampus. Continue removing cortex until you reach the white striations of the corpus callosum. Very slowly and carefully remove the horizontal white striations of the corpus callosum until you reach the striations that go vertical. When you reach these striations, stop aspirating and clean up the sides so that the lens can be placed into the brain. Continue washing with cortex buffer until all bleeding stops and keep the brain constantly flushed with cortex buffer.<br />
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=== Lens implantation ===<br />
Attach GRIN lens to lens holder on stereotax. Align the lens with the top of the skull at the most posterior part of the craniotomy and quickly insert the lens 1.35 mm below the top of the skull. Remove any excess liquid with the aspirator. Connect the outside of the GRIN lens with the skull and skull screw using cyanoacrylate glue and let fully dry or use a glue accelerator. Remove the lens holder by simply removing the vacuum suction (you can just pinch the tubing) and withdrawing the holder. Next, cover the entire skull with cyanoacrylate glue, and then cover with dental cement. Let dry completely and cover with Kwik-Sil to protect the lens. Remove from stereotax and allow the animal to recover. Give the animal amoxicillin (or equivalent) for 7 days, and daily injections of carprofen and dexamethasone for 7 days.<br />
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=== Important tips to remember ===<br />
*Try to keep the surgery as sterile as possible. Infections will destroy imaging and the ability to keep the same cells over many days.<br />
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*Try to keep isoflurane as low as possible. Surgeries can take 2-6 hours depending on experience.<br />
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*Aspirations will take a lot of practice. Go slow at first.<br />
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*Depth will be trial and error. If the first few animals don't give good images, check the depth using histology and adjust accordingly.<br />
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Please feel free to add any tips that you have!<br />
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= Baseplating Protocol =<br />
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== Basic Equipment needed ==<br />
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You will only need basic surgical equipment to baseplate. This includes a mouse stereotax, an isoflurane vaporizer, surgical heat pad, stereo surgical microscope, light source, and a dental drill. <br />
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You will also need cyanoacrylate glue, fine forceps, double distilled water, lens paper, and dental cement<br />
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== Baseplate Preparation ==<br />
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Before affixing the baseplate on an animal, you will need to prepare the baseplate. <br />
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The first step is to insert magnets into the appropriate holes on the baseplate. To ensure that the polarity of the magnets is correct throughout the baseplate preparation, use a reference Miniscope to hold onto the magnets. Mark the top magnet with a sharpie, as seen in the picture below. Next, put the top magnet on the head of a screwdriver with the marked side facing away from the screwdriver. Apply cyanoacrylate glue on the edge of the magnet, then push the magnet through the appropriate hole. Ensure that the magnet goes through the hole completely and the top side of the baseplate is flat, as the quality of imaging relies on the Miniscope sitting flush on the baseplate. Repeat this procedure for the next 2 magnets. Finally, put a coat of cyanoacrylate glue on the bottom side of the baseplate and leave the baseplate to dry.<br />
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Once the baseplate has dried, score the bottom and the sides of the baseplate using a dental drill to allow the dental glue to adhere the baseplate and skull well. <br />
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== Cap Preparation ==<br />
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Similar to the baseplate preparation, use a reference Miniscope to hold onto the magnets. Mark the top magnet with a sharpie, and put the magnet on the head of a screwdriver with the marked side facing toward the screwdriver. Coat the magnet with cyanoacrylate glue and push the magnet into the appropriate hole. Repeat for the remaining two magnets.<br />
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== Baseplating ==<br />
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Connect a Miniscope to a DAQ and initialize the Miniscope software on a computer. Attach your prepared baseplate onto the bottom of the Miniscope using a set screw.<br />
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Using a stereotax and isoflurane vaporizer, anesthetize the animal and fix ear bars until the animal is stable. Carefully remove the Kwik-Sil off the top of the animal’s GRIN lens with blunt forceps being sure not to scratch the lens. Under a stereo surgical microscope, check to see if there is debris on the top of the GRIN lens. If there is, dampen a sheet of lens paper using double distilled water. Wipe the top of the GRIN lens using fine forceps and lens paper, being sure to only wipe laterally and not to apply direct downward pressure.<br />
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After the top of the GRIN lens is clean, explore the field of view with the Miniscope. Adjust the LED brightness and focus accordingly. We advise that you use maximum gain and low LED excitation as to not photobleach the brain during baseplating. After finding your ideal view, practice taking the Miniscope off and on and finding the same view. <br />
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When you feel comfortable repeatedly finding the view with the Miniscope, apply dental cement in a “C” pattern around the GRIN lens, making sure not to touch the lens itself. It will be easier if your first application of dental cement is more viscous. Put the Miniscope on top of the lens and find your ideal view. Hold the Miniscope steadily as the first application of dental cement dries, making sure that your view does not change. <br />
After the first application of dental cement dries, mix more dental cement and apply it on the posterior edge of the baseplate, making sure it seals to the skull. At this point, you can also build up dental cement along the wall of the baseplate on the animal’s right side. Be sure to avoid applying dental cement onto the camera itself and adhering it to the baseplate. Once the dental cement dries and you feel confident that the baseplate will no longer shift, carefully remove the Miniscope from the baseplate by unscrewing the set screw and lifting it off. <br />
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Apply dental cement to the anterior edge of the baseplate and fill in the gaps between the baseplate and skull on the animal’s left side. Allow the dental cement to dry and attach the plastic cap onto the top of the baseplate with a set screw.</div>Chrisl1026