GI - User contributions [en]

Nikon TIRF switch on

2026-06-10T09:12:38Z

Jelle: reordered

[[File:TIRF switch on.png|880x880px]]

=== '''Required elements''' ===
4) '''Controller''' (this operates the microscope)

5) '''Internal camera''' (Fi3, to inspect the objective back aperture to guarantee TIRF, software always requires this on)

6) '''Main camera''' (Prime BSI, to actually observe the sample through the objective lens)

7) '''Computer''' (for observation and imaging by camera, otherwise you can only use the eyepieces)

=== '''Optional elements''' ===
1) '''Lasers''' (if you want to actually do TIRF, or use the specific laser wavelengths for widefield epifluorescence)

2) '''Warming box''' (if you want to have the incubator box light available, and for if you want to do live-cell incubation)

3) '''Humidity + CO2''' (for live-cell incubation)

'''-> When in doubt:''' Switch them all on

Nikon TIRF widefield

2026-06-10T09:11:39Z

Jelle:

== Manuals and user guides ==
[[TIRF alignment guide]]

[[Nikon TIRF switch on|How to switch on]]
== Current status ==
The current status of the microscope can be found [[Current microscope status#Light microscopes|here]] and on the [https://bookings.science.ru.nl/category/general-instrumentation-gi/gi-light-microscopy-image-analysis booking website].
== Biosafety and access ==
The TIRF is located in an ML-1 room. See the [[General Instrumentation Department#Biosafety|Biosafety]] section on rules regarding working in this room, and how to get access.

== Technical specifications ==
'''Light sources and filters:'''

'''Epifluorescence mode''' with LED light sources (''non-TIRF)'': Excitations: 385, 475, 550, 621 nm. Emissions: Band pass emission filters for blue (ca. 450), green (ca. 510), red (ca. 600) and far-red (ca. 670).

'''TIRF-mode''' with lasers: Excitations: 488 nm, 561 nm, 640 nm. Emission filters: None

Nikon TIRF switch on

2026-06-10T09:11:11Z

Jelle: completed the steps

[[File:TIRF switch on.png|880x880px]]

=== '''Required elements''' ===
4) '''Controller''' (this operates the microscope)

5) '''Internal camera''' (Fi3, to inspect the objective back aperture to guarantee TIRF, software always requires this on)

6) '''Main camera''' (Prime BSI, to actually observe the sample through the objective lens)

=== '''Optional elements''' ===
1) '''Lasers''' (if you want to actually do TIRF, or use the specific laser wavelengths for widefield epifluorescence)

2) '''Warming box''' (if you want to have the incubator box light available, and for if you want to do live-cell incubation)

3) '''Humidity + CO2''' (for live-cell incubation)

7) '''Computer''' (for observation and imaging by camera, otherwise you can only use the eyepieces)

'''-> When in doubt:''' Switch them all on

Nikon TIRF switch on

2026-06-10T08:03:35Z

Jelle: dsf

[[File:TIRF switch on.png|880x880px]]

Required elements

4: Controller (this operates the microscope)

5: Internal cam (to inspect the objective back aperture to guarantee TIRF, software always requires this on)

Optional elements

1: Lasers (if you want to do TIRF or use lasers for widefield)

2: Warming box (if you want to have the box-light available or for live-cell incubation)

3: CO2 and humidity (for live-cell incubation)

File:TIRF switch on.png

2026-06-10T08:00:52Z

Jelle:

which components to switch on to switch on the TIRF microscope and in which order

Nikon TIRF widefield

2026-06-10T07:59:47Z

Jelle:

== Manuals and user guides ==
[[TIRF alignment guide]]

How to switch on
== Current status ==
The current status of the microscope can be found [[Current microscope status#Light microscopes|here]] and on the [https://bookings.science.ru.nl/category/general-instrumentation-gi/gi-light-microscopy-image-analysis booking website].
== Biosafety and access ==
The TIRF is located in an ML-1 room. See the [[General Instrumentation Department#Biosafety|Biosafety]] section on rules regarding working in this room, and how to get access.

== Technical specifications ==
'''Light sources and filters:'''

'''Epifluorescence mode''' with LED light sources (''non-TIRF)'': Excitations: 385, 475, 550, 621 nm. Emissions: Band pass emission filters for blue (ca. 450), green (ca. 510), red (ca. 600) and far-red (ca. 670).

'''TIRF-mode''' with lasers: Excitations: 488 nm, 561 nm, 640 nm. Emission filters: None

Nikon sora introduction protocol

2026-05-28T15:40:07Z

Jelle:

=== '''Introduction protocol''' ===

'''Type of microscope and its strengths'''

* Spinning disc microscope for fast confocal microscopy (optical sectioning)
* Suitable for live-cell imaging experiments
* Superresolution optional, to achieve (ideally) ca. 150 nm resolution in green.
* Two-camera option for capturing two fluorescent colors at the same time (dynamic processes)
* Fluorescence Recovery After Photobleaching (FRAP) to study molecular diffusion times

[[Nikon sora switch on|'''Switch-on procedure''']] (link)

* Make sure to also start the software already and start Single-cam mode

'''Stage-top incubation system for live-cell imaging (switch this on before starting a live cell experiment)'''

* Component 2 is the "Warming box" and needs to be switched on as well as the "Heater" button on its front panel
* Component 3 needs to be switched on and is already on the right settings.
* Look at the stage-top incubator itself - it has a lid that should be used during the experiment
* There is a plastic removable frame covering a "canal" (basin) of water around the edge of the sample area
* The "canal" should be filled with MQ water using the attached tubing - the tubing sticks out of the front of the microscope
* Unclip the tubing clip, attach a syringe with MQ water fill stage-top incubator "canal" with MQ water
* Clip the tubing clip again, remove the syringe, and cover the canal with the plastic frame. It is ready to use<br />
'''First manual operations (moving things around)'''

* Operate the lens switcher at the front of the microscope and select the 100X / Oil lens.
* Operate the joystick in various directions as an example. You may have to switch its movement speed to fast.
* Center the stage above the objective.
* Operate the focus knob on the side of the joystick to see the objective move up and down (also may have to switch the movement speed to fast).
* Lower the lens as far as possible down to prepare for placing the sample.<br />
'''First brightfield observation (finding the sample)'''

* Mount a demo sample (or user sample) in the appropriate sample holder without placing it in the microscope yet
* Put oil on the lens
* Put the condenser arm in the backward position (access to sample area easier)
* Mount the sample holder with sample onto the microscope stage
* Make sure the condenser arm (top arm of the microscope) is set to "O" (brightfield shutter = Open) and not "C"
* Restore the condenser arm to the normal operating position (upright, not backwards)
* Switch on the brightfield lamp (lamp icon, left side of the microscope body) and demonstrate adjusting light intensity (left side rotatable adjuster knob)
* Make sure the eye-piece is active using the "eye" button at the front of the microscope
* Confirm with the user that light is visible through the microscope and that it responds to intensity adjustment
* Visually point the brightfield-lamp-spot onto a relevant sample area, where we expect to see structures
* Let the user raise the objective while looking through the eyepieces, until the sample becomes visible

'''Epifluorescence mode by camera (quick checking fluorescent signals)'''

* Go to the computer and make sure everyone can see the screen
* Point out that the software looks complicated but we will discuss the essential parts to understand them
* Point out that there are modalities and buttons to switch to them (Eye-observation, Epifluorescence with camera, Spinning-disc modalities).
* Point out that we were using the Eye-observation mode all this time
* Switch to Epifluorescence with camera mode
* Choose a pre-set optical configuration that should reveal a fluorescent color that is in the sample
* Point out that the two settings that matter now are a light intensity slider (lower left) and a camera exposure time (top left), set them both to low values
* Click Capture image and probably see nothing (black image, but it may just be a low amount of signal!)
* Switch on auto-scale (above the picture, a graph symbol with a Play button on top of it) and explain that this will always show what the camera sees, even if it is weak or strong signal. Now you will see something.
* Click Live to live-view with the camera for making further adjustments
* Raise the light source and exposure times and see the signal becoming higher than the noise, explain why/that it should be higher than the noise
* Then lower the both down and see the signal disappearing in the noise (undesirable in a real experiment)
* Save the light and exposure time adjustments using right-click on the Optical Configuration and choosing "Assign" - explain that this is necessary upon changing something
* Select another pre-set (e.g. mCherry) and point out that another fluorescent color will be activated now and why
* Move on from Epifluorescence-by-eye mode as this is not worth spending the users' brain power on too much.

'''Confocal Spinning Disc mode (the main thing you do with this microscope)'''

* Switch to the Spinning Disc modality
* Activate a pre-set, e.g. "488"
* Point out that it now looks different but we will also go step-by-step
* Point out that there are now 5 lasers available
* Capture Image and inspect the result
* Live view and adjust laser and exposure time to get decent signal
* Point out that we are now in confocal mode and the optical section is thinner than before
* Let the user move the focus up/down and see clearly different layers of the sample, especially noticable because it's in confocal mode
* Point out the lack of out-of-focus blur to highlight the confocality again.
* Adjust the light intensity and exposure times to show that the same principles apply, you need signals stronger than noise.
* Repeat this for another pre-set Optical Configuration.<br />
'''Camera settings in more detail (worth spending time on this)'''

* ''Binning'': You can choose to sacrifice resolution to gain sensitivity to low signals by increasing binning
* ''Bit-depth'': Always 16-bit, because it has the most accurate intensity registration. 12-bit doesn't make sense to use for any obvious reason.
* ''Exposure time'': Adjust in multiples of 5. You can show what happens with an exposure time of 2 ms (stripes!)
* ''Scan mode'': Always on "Fast" (the other ones are just doing on-chip averaging to get more signal-to-noise, whereas you can just increase exposure time to achieve the same).
* ''Full-sensor or smaller sensor area (custom)'': Start with full-sensor always. The only application for a smaller sensor size is when you need extremely rapid imaging, as those images will then be smaller in data-size to transfer from the camera to the computer.

'''[[Nikon sora superresolution switch|SoRa Superresolution, how to switch on/off (link)]]'''

* This should theoretically be able to reach resolutions of around 150 nm (at 510 nm)

'''ND Acquisition'''

This is used to do automated multidimensional experiments

Start with a Z-stack, because the full SoRa pipeline to get maximum resolution needs this.

Perfect Focus System for keeping/using exact focus levels in live view and during ND Acquisition experiments

FRAP (optional)

Dual camera mode (optional)

Data saving (nd2 files) and transfering (SURF file sender)

User account creation

Booking system

Current microscope status

2026-05-28T14:48:06Z

Jelle: /* Light microscopes */

=== Light microscopes ===
{| class="wikitable"
|+
!Microscope
!Status
!Notes
!Date
|-
|[[Leica SP8X|SP8X]]
|OK
|
|
|-
|[[Leica SP8 Liachroic|SP8-Lia]]
|OK
|488 laser functional again
|20.11.2025
|-
|[[Leica Thunder|Thunder]]
|OK
|
|
|-
|[[Leica Timelapse|Timelapse]]
|OK
|New camera installed
|25.09.2025
|-
|[[PhaseView light sheet|Lightsheet]]
|OK
|Microscope fully functional, use 4TB SSD drive to take images
|20.11.2025
|-
|[[DM2500]]
|OK
|
|
|-
|[[Nikon sora|SoRa]]
|OK
|
|
|-
|[[Nikon TIRF widefield|TIRF]]
|OK
|
|
|-
|Spinoza server
|OK
|
|
|}

=== Electron microscopes and sample prep equipment ===
{| class="wikitable"
|+
!Machine
!Status
!Notes
!Date
|-
|[[JEOL1400 TEM|JEOL1400]]
|OK
|
|
|-
|[[JEOL2100 TEM|JEOL2100]]
|OK
|HT repaired feb 24-26

|11.03.2026
|-
|
|
|STEM detector broken - under repair
EDX detector broken
|aug 2024
pre 2023
|-
|[[Cryo-holder, cryo-dock and cryo-pumping station|Cryo-dock/station/holder]]
|OK
|
|
|-
|[[HPM100]]
|OK
|
|
|-
|[[EMPACT|EMpact]]
|Broken
|Unsure if can be fixed
|01.09.2025
|-
|[[Vitrobot]]
|OK
|
|
|-
|[[UC-E microtome]]
|OK
|
|
|-
|UC7/FC7 cryo-microtome
|OK
|
|
|-
|[[CCU-010]]
|OK
|
|
|-
|Edward306
|Broken
|Under repair
|
|-
|BAF400
|OK
|
|
|-
|Vibratome
|OK
|
|
|-
|CPD-020
|In storage
|
|
|-
|JFD-030
|OK
|
|
|}

Nikon sora introduction protocol

2026-05-28T07:57:50Z

Jelle:

=== '''Introduction protocol''' ===

'''Type of microscope and its strengths'''

* Spinning disc microscope for fast confocal microscopy (optical sectioning)
* Suitable for live-cell imaging experiments
* Superresolution optional, to achieve (ideally) ca. 150 nm resolution in green.
* Two-camera option for capturing two fluorescent colors at the same time (dynamic processes)
* Fluorescence Recovery After Photobleaching (FRAP) to study molecular diffusion times

[[Nikon sora switch on|'''Switch-on procedure''']] (link)

* Make sure to also start the software already and start Single-cam mode

'''Stage-top incubation system for live-cell imaging (switch on before starting a live cell experiment)'''

* Component 2 is the "Warming box" and needs to be switched on as well as the "Heater" button on its front panel
* Component 3 needs to be switched on and is already on the right settings.
* Look at the stage-top incubator itself - it has a lid that should be used during the experiment
* There is a plastic removable frame covering a "canal" (basin) of water around the edge of the sample area
* The "canal" should be filled with MQ water using the attached tubing - the tubing sticks out of the front of the microscope
* Unclip the tubing clip, attach a syringe with MQ water fill stage-top incubator "canal" with MQ water
* Clip the tubing clip again, remove the syringe, and cover the canal with the plastic frame. It is ready to use

'''First manual operations'''

* Operate the lens switcher at the front of the microscope and select the 100X / Oil lens.
* Operate the joystick in various directions as an example. You may have to switch its movement speed to fast.
* Center the stage above the objective.
* Operate the focus knob on the side of the joystick to see the objective move up and down (also may have to switch the movement speed to fast).
* Lower the lens as far as possible down to prepare for placing the sample.

'''First brightfield observation'''

* Mount a demo sample (or user sample) in the appropriate sample holder without placing it in the microscope yet
* Put oil on the lens
* Put the condenser arm in the backward position (access to sample area easier)
* Mount the sample holder with sample onto the microscope stage
* Make sure the condenser arm (top arm of the microscope) is set to "O" (brightfield shutter = Open) and not "C"
* Restore the condenser arm to the normal operating position (upright, not backwards)
* Switch on the brightfield lamp (lamp icon, left side of the microscope body) and demonstrate adjusting light intensity (left side rotatable adjuster knob)
* Make sure the eye-piece is active using the "eye" button at the front of the microscope
* Confirm with the user that light is visible through the microscope and that it responds to intensity adjustment
* Visually point the brightfield-lamp-spot onto a relevant sample area, where we expect to see structures
* Let the user raise the objective while looking through the eyepieces, until the sample becomes visible

'''Epifluorescence mode by camera (quick checking fluorescent signals)'''

* Go to the computer and make sure everyone can see the screen
* Point out that the software looks complicated but we will discuss the essential parts to understand them
* Roughly describe that there are modalities (Eye-observation, Epifluorescence with camera, Spinning-disc modalities).
* Point out that we were using the Eye-observation mode all this time
* Switch to Epifluorescence with camera mode
* Choose a pre-set optical configuration that should reveal a fluorescent color that is in the sample
* Point out that the two settings that matter now are a light intensity slider (lower left) and a camera exposure time (top left), set them both to low values
* Click Capture image and probably see nothing (black image, but it may just be a low amount of signal!)
* Switch on auto-scale (above the picture, a graph symbol with a Play button on top of it) and explain that this will always show what the camera sees, even if it is weak or strong signal. Now you will see something.
* Click Live to live-view with the camera for making further adjustments
* Raise the light source and exposure times and see the signal becoming higher than the noise, explain why/that it should be higher than the noise
* Then lower the both down and see the signal disappearing in the noise (undesirable in a real experiment)
* Save the light and exposure time adjustments using right-click on the Optical Configuration and choosing "Assign" - explain that this is necessary upon changing something
* Select another pre-set (e.g. mCherry) and point out that another fluorescent color will be activated now and why
* Move on from Epifluorescence-by-eye mode as this is not worth spending the users' brain power on too much.

'''Confocal Spinning Disc mode'''

* Switch to the Spinning Disc modality
* Activate a pre-set, e.g. "488"
* Point out that it now looks different but we will also go step-by-step
* Point out that there are now 5 lasers available
* Capture Image and inspect the result
* Live view and adjust laser and exposure time to get decent signal
* Point out that we are now in confocal mode and the optical section is thinner than before
* Let the user move the focus up/down and see clearly different layers of the sample, especially noticable because it's in confocal mode
* Point out the lack of out-of-focus blur to highlight the confocality again.
* Adjust the light intensity and exposure times to show that the same principles apply, you need signals stronger than noise.
* Repeat this for another pre-set Optical Configuration.

'''In-depth camera settings (worth spending time on)'''

* ''Binning'': You can choose to sacrifice resolution to gain sensitivity to low signals by increasing binning
* ''Bit-depth'': Always 16-bit, because it has the most accurate intensity registration. 12-bit doesn't make sense to use for any obvious reason.
* ''Exposure time'': Adjust in multiples of 5. You can show what happens with an exposure time of 2 ms (stripes!)
* ''Scan mode'': Always on "Fast" (the other ones are just doing on-chip averaging to get more signal-to-noise, whereas you can just increase exposure time to achieve the same).
* ''Full-sensor or smaller sensor area (custom)'': Start with full-sensor always. The only application for a smaller sensor size is when you need extremely rapid imaging, as those images will then be smaller in data-size to transfer from the camera to the computer.

'''[[Nikon sora superresolution switch|SoRa Superresolution, how to switch on/off (link)]]'''

* This should theoretically reach resolutions of ca. 150 nm (green)

ND Acquisition for automated multidimensional experiments, start with Z-stack because full SoRa pipeline needs this.

Perfect Focus System for keeping/using exact focus levels in live view and during ND Acquisition experiments

FRAP (optional)

Dual camera mode (optional)

Data saving (nd2 files) and transfering (SURF file sender)

User account creation

Booking system

Nikon sora introduction protocol

2026-05-28T07:56:25Z

Jelle:

=== '''Introduction protocol''' ===

'''Type of microscope and its strengths'''

* Spinning disc microscope for fast confocal microscopy (optical sectioning)
* Suitable for live-cell imaging experiments
* Superresolution optional, to achieve (ideally) ca. 150 nm resolution in green.
* Two-camera option for capturing two fluorescent colors at the same time (dynamic processes)
* Fluorescence Recovery After Photobleaching (FRAP) to study molecular diffusion times

[[Nikon sora switch on|'''Switch-on procedure''']]

* Make sure to also start the software already and start Single-cam mode

'''Stage-top incubation system for live-cell imaging (switch on before starting a live cell experiment)'''

* Component 2 is the "Warming box" and needs to be switched on as well as the "Heater" button on its front panel
* Component 3 needs to be switched on and is already on the right settings.
* Look at the stage-top incubator itself - it has a lid that should be used during the experiment
* There is a plastic removable frame covering a "canal" (basin) of water around the edge of the sample area
* The "canal" should be filled with MQ water using the attached tubing - the tubing sticks out of the front of the microscope
* Unclip the tubing clip, attach a syringe with MQ water fill stage-top incubator "canal" with MQ water
* Clip the tubing clip again, remove the syringe, and cover the canal with the plastic frame. It is ready to use

'''First manual operations'''

* Operate the lens switcher at the front of the microscope and select the 100X / Oil lens.
* Operate the joystick in various directions as an example. You may have to switch its movement speed to fast.
* Center the stage above the objective.
* Operate the focus knob on the side of the joystick to see the objective move up and down (also may have to switch the movement speed to fast).
* Lower the lens as far as possible down to prepare for placing the sample.

'''First brightfield observation'''

* Mount a demo sample (or user sample) in the appropriate sample holder without placing it in the microscope yet
* Put oil on the lens
* Put the condenser arm in the backward position (access to sample area easier)
* Mount the sample holder with sample onto the microscope stage
* Make sure the condenser arm (top arm of the microscope) is set to "O" (brightfield shutter = Open) and not "C"
* Restore the condenser arm to the normal operating position (upright, not backwards)
* Switch on the brightfield lamp (lamp icon, left side of the microscope body) and demonstrate adjusting light intensity (left side rotatable adjuster knob)
* Make sure the eye-piece is active using the "eye" button at the front of the microscope
* Confirm with the user that light is visible through the microscope and that it responds to intensity adjustment
* Visually point the brightfield-lamp-spot onto a relevant sample area, where we expect to see structures
* Let the user raise the objective while looking through the eyepieces, until the sample becomes visible

'''Epifluorescence mode by camera (quick checking fluorescent signals)'''

* Go to the computer and make sure everyone can see the screen
* Point out that the software looks complicated but we will discuss the essential parts to understand them
* Roughly describe that there are modalities (Eye-observation, Epifluorescence with camera, Spinning-disc modalities).
* Point out that we were using the Eye-observation mode all this time
* Switch to Epifluorescence with camera mode
* Choose a pre-set optical configuration that should reveal a fluorescent color that is in the sample
* Point out that the two settings that matter now are a light intensity slider (lower left) and a camera exposure time (top left), set them both to low values
* Click Capture image and probably see nothing (black image, but it may just be a low amount of signal!)
* Switch on auto-scale (above the picture, a graph symbol with a Play button on top of it) and explain that this will always show what the camera sees, even if it is weak or strong signal. Now you will see something.
* Click Live to live-view with the camera for making further adjustments
* Raise the light source and exposure times and see the signal becoming higher than the noise, explain why/that it should be higher than the noise
* Then lower the both down and see the signal disappearing in the noise (undesirable in a real experiment)
* Save the light and exposure time adjustments using right-click on the Optical Configuration and choosing "Assign" - explain that this is necessary upon changing something
* Select another pre-set (e.g. mCherry) and point out that another fluorescent color will be activated now and why
* Move on from Epifluorescence-by-eye mode as this is not worth spending the users' brain power on too much.

'''Confocal Spinning Disc mode'''

* Switch to the Spinning Disc modality
* Activate a pre-set, e.g. "488"
* Point out that it now looks different but we will also go step-by-step
* Point out that there are now 5 lasers available
* Capture Image and inspect the result
* Live view and adjust laser and exposure time to get decent signal
* Point out that we are now in confocal mode and the optical section is thinner than before
* Let the user move the focus up/down and see clearly different layers of the sample, especially noticable because it's in confocal mode
* Point out the lack of out-of-focus blur to highlight the confocality again.
* Adjust the light intensity and exposure times to show that the same principles apply, you need signals stronger than noise.
* Repeat this for another pre-set Optical Configuration.

'''In-depth camera settings (worth spending time on)'''

* ''Binning'': You can choose to sacrifice resolution to gain sensitivity to low signals by increasing binning
* ''Bit-depth'': Always 16-bit, because it has the most accurate intensity registration. 12-bit doesn't make sense to use for any obvious reason.
* ''Exposure time'': Adjust in multiples of 5. You can show what happens with an exposure time of 2 ms (stripes!)
* ''Scan mode'': Always on "Fast" (the other ones are just doing on-chip averaging to get more signal-to-noise, whereas you can just increase exposure time to achieve the same).
* ''Full-sensor or smaller sensor area (custom)'': Start with full-sensor always. The only application for a smaller sensor size is when you need extremely rapid imaging, as those images will then be smaller in data-size to transfer from the camera to the computer.

SoRa Superresolution to double resolving power

ND Acquisition for automated multidimensional experiments, start with Z-stack because full SoRa pipeline needs this.

Perfect Focus System for keeping/using exact focus levels in live view and during ND Acquisition experiments

FRAP (optional)

Dual camera mode (optional)

Data saving (nd2 files) and transfering (SURF file sender)

User account creation

Booking system

Nikon sora introduction protocol

2026-05-27T16:34:40Z

Jelle:

=== '''Introduction protocol''' ===

'''Type of microscope and its strengths'''

* Spinning disc microscope for fast confocal microscopy (optical sectioning)
* Suitable for live-cell imaging experiments
* Superresolution optional, to achieve (ideally) ca. 150 nm resolution in green.
* Two-camera option for capturing two fluorescent colors at the same time (dynamic processes)
* Fluorescence Recovery After Photobleaching (FRAP) to study molecular diffusion times

[[Nikon sora switch on|'''Switch-on procedure''']]

* Make sure to also start the software already and start Single-cam mode

'''Stage-top incubation system for live-cell imaging (switch on before starting a live cell experiment)'''

* Component 2 is the "Warming box" and needs to be switched on as well as the "Heater" button on its front panel
* Look at the stage-top incubator itself - it has a lid that should be used during the experiment
* There is a plastic removable frame covering a "canal" (basin) of water around the edge of the sample area
* The "canal" should be filled with MQ water using the attached tubing - the tubing sticks out of the front of the microscope
* Unclip the tubing clip, attach a syringe with MQ water fill stage-top incubator "canal" with MQ water
* Clip the tubing clip again, remove the syringe, and cover the canal with the plastic frame. It is ready to use

'''First manual operations'''

* Operate the lens switcher at the front of the microscope and select the 100X / Oil lens.
* Operate the joystick in various directions as an example. You may have to switch its movement speed to fast.
* Center the stage above the objective.
* Operate the focus knob on the side of the joystick to see the objective move up and down (also may have to switch the movement speed to fast).
* Lower the lens as far as possible down to prepare for placing the sample.

'''First brightfield observation'''

* Mount a demo sample (or user sample) in the appropriate sample holder without placing it in the microscope yet
* Put oil on the lens
* Put the condenser arm in the backward position (access to sample area easier)
* Mount the sample holder with sample onto the microscope stage
* Make sure the condenser arm (top arm of the microscope) is set to "O" (brightfield shutter = Open) and not "C"
* Restore the condenser arm to the normal operating position (upright, not backwards)
* Switch on the brightfield lamp (lamp icon, left side of the microscope body) and demonstrate adjusting light intensity (left side rotatable adjuster knob)
* Make sure the eye-piece is active using the "eye" button at the front of the microscope
* Confirm with the user that light is visible through the microscope and that it responds to intensity adjustment
* Visually point the brightfield-lamp-spot onto a relevant sample area, where we expect to see structures
* Let the user raise the objective while looking through the eyepieces, until the sample becomes visible

'''Epifluorescence mode by camera (quick checking fluorescent signals)'''

* Go to the computer and make sure everyone can see the screen
* Point out that the software looks complicated but we will discuss the essential parts to understand them
* Roughly describe that there are modalities (Eye-observation, Epifluorescence with camera, Spinning-disc modalities).
* Point out that we were using the Eye-observation mode all this time
* Switch to Epifluorescence with camera mode
* Choose a pre-set optical configuration that should reveal a fluorescent color that is in the sample
* Point out that the two settings that matter now are a light intensity slider (lower left) and a camera exposure time (top left), set them both to low values
* Click Capture image and probably see nothing (black image, but it may just be a low amount of signal!)
* Switch on auto-scale (above the picture, a graph symbol with a Play button on top of it) and explain that this will always show what the camera sees, even if it is weak or strong signal. Now you will see something.
* Click Live to live-view with the camera for making further adjustments
* Raise the light source and exposure times and see the signal becoming higher than the noise, explain why/that it should be higher than the noise
* Then lower the both down and see the signal disappearing in the noise (undesirable in a real experiment)
* Save the light and exposure time adjustments using right-click on the Optical Configuration and choosing "Assign" - explain that this is necessary upon changing something
* Select another pre-set (e.g. mCherry) and point out that another fluorescent color will be activated now and why
* Move on from Epifluorescence-by-eye mode as this is not worth spending the users' brain power on too much.

'''Confocal Spinning Disc mode'''

* Switch to the Spinning Disc modality
* Activate a pre-set, e.g. "488"
* Point out that it now looks different but we will also go step-by-step
* Point out that there are now 5 lasers available
* Capture Image and inspect the result
* Live view and adjust laser and exposure time to get decent signal
* Point out that we are now in confocal mode and the optical section is thinner than before
* Let the user move the focus up/down and see clearly different layers of the sample, especially noticable because it's in confocal mode
* Point out the lack of out-of-focus blur to highlight the confocality again.
* Adjust the light intensity and exposure times to show that the same principles apply, you need signals stronger than noise.
* Repeat this for another pre-set Optical Configuration.

'''In-depth camera settings (worth spending time on)'''

* ''Binning'': You can choose to sacrifice resolution to gain sensitivity to low signals by increasing binning
* ''Bit-depth'': Always 16-bit, because it has the most accurate intensity registration. 12-bit doesn't make sense to use for any obvious reason.
* ''Exposure time'': Adjust in multiples of 5. You can show what happens with an exposure time of 2 ms (stripes!)
* ''Scan mode'': Always on "Fast" (the other ones are just doing on-chip averaging to get more signal-to-noise, whereas you can just increase exposure time to achieve the same).
* ''Full-sensor or smaller sensor area (custom)'': Start with full-sensor always. The only application for a smaller sensor size is when you need extremely rapid imaging, as those images will then be smaller in data-size to transfer from the camera to the computer.

SoRa Superresolution to double resolving power

ND Acquisition for automated multidimensional experiments, start with Z-stack because full SoRa pipeline needs this.

Perfect Focus System for keeping/using exact focus levels in live view and during ND Acquisition experiments

FRAP (optional)

Dual camera mode (optional)

Data saving (nd2 files) and transfering (SURF file sender)

User account creation

Booking system

Nikon sora introduction protocol

2026-05-27T16:34:14Z

Jelle:

=== '''Introduction protocol''' ===

'''Type of microscope and its strengths'''

* Spinning disc microscope for fast confocal microscopy (optical sectioning)
* Suitable for live-cell imaging experiments
* Superresolution optional, to achieve (ideally) ca. 150 nm resolution in green.
* Two-camera option for capturing two fluorescent colors at the same time (dynamic processes)
* Fluorescence Recovery After Photobleaching (FRAP) to study molecular diffusion times
[[Nikon sora switch on|Switch-on procedure]]

[[Nikon sora switch on|'''Switch-on procedure''']]

* Make sure to also start the software already and start Single-cam mode

'''Stage-top incubation system for live-cell imaging (switch on before starting a live cell experiment)'''

* Component 2 is the "Warming box" and needs to be switched on as well as the "Heater" button on its front panel
* Look at the stage-top incubator itself - it has a lid that should be used during the experiment
* There is a plastic removable frame covering a "canal" (basin) of water around the edge of the sample area
* The "canal" should be filled with MQ water using the attached tubing - the tubing sticks out of the front of the microscope
* Unclip the tubing clip, attach a syringe with MQ water fill stage-top incubator "canal" with MQ water
* Clip the tubing clip again, remove the syringe, and cover the canal with the plastic frame. It is ready to use
First manual operation

'''First manual operations'''

* Operate the lens switcher at the front of the microscope and select the 100X / Oil lens.
* Operate the joystick in various directions as an example. You may have to switch its movement speed to fast.
* Center the stage above the objective.
* Operate the focus knob on the side of the joystick to see the objective move up and down (also may have to switch the movement speed to fast).
* Lower the lens as far as possible down to prepare for placing the sample.
First brightfield operation

'''First brightfield observation'''

* Mount a demo sample (or user sample) in the appropriate sample holder without placing it in the microscope yet
* Put oil on the lens
* Put the condenser arm in the backward position (access to sample area easier)
* Mount the sample holder with sample onto the microscope stage
* Make sure the condenser arm (top arm of the microscope) is set to "O" (brightfield shutter = Open) and not "C"
* Restore the condenser arm to the normal operating position (upright, not backwards)
* Switch on the brightfield lamp (lamp icon, left side of the microscope body) and demonstrate adjusting light intensity (left side rotatable adjuster knob)
* Make sure the eye-piece is active using the "eye" button at the front of the microscope
* Confirm with the user that light is visible through the microscope and that it responds to intensity adjustment
* Visually point the brightfield-lamp-spot onto a relevant sample area, where we expect to see structures
* Let the user raise the objective while looking through the eyepieces, until the sample becomes visible

'''Epifluorescence mode by camera (quick checking fluorescent signals)'''

* Go to the computer and make sure everyone can see the screen
* Point out that the software looks complicated but we will discuss the essential parts to understand them
* Roughly describe that there are modalities (Eye-observation, Epifluorescence with camera, Spinning-disc modalities).
* Point out that we were using the Eye-observation mode all this time
* Switch to Epifluorescence with camera mode
* Choose a pre-set optical configuration that should reveal a fluorescent color that is in the sample
* Point out that the two settings that matter now are a light intensity slider (lower left) and a camera exposure time (top left), set them both to low values
* Click Capture image and probably see nothing (black image, but it may just be a low amount of signal!)
* Switch on auto-scale (above the picture, a graph symbol with a Play button on top of it) and explain that this will always show what the camera sees, even if it is weak or strong signal. Now you will see something.
* Click Live to live-view with the camera for making further adjustments
* Raise the light source and exposure times and see the signal becoming higher than the noise, explain why/that it should be higher than the noise
* Then lower the both down and see the signal disappearing in the noise (undesirable in a real experiment)
* Save the light and exposure time adjustments using right-click on the Optical Configuration and choosing "Assign" - explain that this is necessary upon changing something
* Select another pre-set (e.g. mCherry) and point out that another fluorescent color will be activated now and why
* Move on from Epifluorescence-by-eye mode as this is not worth spending the users' brain power on too much.

'''Confocal Spinning Disc mode'''

* Switch to the Spinning Disc modality
* Activate a pre-set, e.g. "488"
* Point out that it now looks different but we will also go step-by-step
* Point out that there are now 5 lasers available
* Capture Image and inspect the result
* Live view and adjust laser and exposure time to get decent signal
* Point out that we are now in confocal mode and the optical section is thinner than before
* Let the user move the focus up/down and see clearly different layers of the sample, especially noticable because it's in confocal mode
* Point out the lack of out-of-focus blur to highlight the confocality again.
* Adjust the light intensity and exposure times to show that the same principles apply, you need signals stronger than noise.
* Repeat this for another pre-set Optical Configuration.

'''In-depth camera settings (worth spending time on)'''

* ''Binning'': You can choose to sacrifice resolution to gain sensitivity to low signals by increasing binning
* ''Bit-depth'': Always 16-bit, because it has the most accurate intensity registration. 12-bit doesn't make sense to use for any obvious reason.
* ''Exposure time'': Adjust in multiples of 5. You can show what happens with an exposure time of 2 ms (stripes!)
* ''Scan mode'': Always on "Fast" (the other ones are just doing on-chip averaging to get more signal-to-noise, whereas you can just increase exposure time to achieve the same).
* ''Full-sensor or smaller sensor area (custom)'': Start with full-sensor always. The only application for a smaller sensor size is when you need extremely rapid imaging, as those images will then be smaller in data-size to transfer from the camera to the computer.

SoRa Superresolution to double resolving power

ND Acquisition for automated multidimensional experiments, start with Z-stack because full SoRa pipeline needs this.

Perfect Focus System for keeping/using exact focus levels in live view and during ND Acquisition experiments

FRAP (optional)

Dual camera mode (optional)

Data saving (nd2 files) and transfering (SURF file sender)

User account creation

Booking system

Nikon sora introduction protocol

2026-05-27T16:27:10Z

Jelle:

'''Introduction protocol'''

Type of microscope and its strengths

* Spinning disc microscope for fast confocal microscopy (optical sectioning)
* Suitable for live-cell imaging experiments
* Superresolution optional, to achieve (ideally) ca. 150 nm resolution in green.
* Two-camera option for capturing two fluorescent colors at the same time (dynamic processes)
* Fluorescence Recovery After Photobleaching (FRAP) to study molecular diffusion times

[[Nikon sora switch on|Switch-on procedure]]

* Make sure to also start the software already and start Single-cam mode
Stage-top incubation system for live-cell imaging (switch on before starting a live cell experiment)

* Component 2 is the "Warming box" and needs to be switched on as well as the "Heater" button on its front panel
* Look at the stage-top incubator itself - it has a lid that should be used during the experiment
* There is a plastic removable frame covering a "canal" (basin) of water around the edge of the sample area
* The "canal" should be filled with MQ water using the attached tubing - the tubing sticks out of the front of the microscope
* Unclip the tubing clip, attach a syringe with MQ water fill stage-top incubator "canal" with MQ water
* Clip the tubing clip again, remove the syringe, and cover the canal with the plastic frame. It is ready to use

First manual operation

* Operate the lens switcher at the front of the microscope and select the 100X / Oil lens.
* Operate the joystick in various directions as an example. You may have to switch its movement speed to fast.
* Center the stage above the objective.
* Operate the focus knob on the side of the joystick to see the objective move up and down (also may have to switch the movement speed to fast).
* Lower the lens as far as possible down to prepare for placing the sample.

First brightfield operation

* Mount a demo sample (or user sample) in the appropriate sample holder without placing it in the microscope yet
* Put oil on the lens
* Put the condenser arm in the backward position (access to sample area easier)
* Mount the sample holder with sample onto the microscope stage
* Make sure the condenser arm (top arm of the microscope) is set to "O" (brightfield shutter = Open) and not "C"
* Restore the condenser arm to the normal operating position (upright, not backwards)
* Switch on the brightfield lamp (lamp icon, left side of the microscope body) and demonstrate adjusting light intensity (left side rotatable adjuster knob)
* Make sure the eye-piece is active using the "eye" button at the front of the microscope
* Confirm with the user that light is visible through the microscope and that it responds to intensity adjustment
* Visually point the brightfield-lamp-spot onto a relevant sample area, where we expect to see structures
* Let the user raise the objective while looking through the eyepieces, until the sample becomes visible

Epifluorescence mode by camera (quick checking fluorescent signals)

* Go to the computer and make sure everyone can see the screen
* Point out that the software looks complicated but we will discuss the essential parts to understand them
* Roughly describe that there are modalities (Eye-observation, Epifluorescence with camera, Spinning-disc modalities).
* Point out that we were using the Eye-observation mode all this time
* Switch to Epifluorescence with camera mode
* Choose a pre-set optical configuration that should reveal a fluorescent color that is in the sample
* Point out that the two settings that matter now are a light intensity slider (lower left) and a camera exposure time (top left), set them both to low values
* Click Capture image and probably see nothing (black image, but it may just be a low amount of signal!)
* Switch on auto-scale (above the picture, a graph symbol with a Play button on top of it) and explain that this will always show what the camera sees, even if it is weak or strong signal. Now you will see something.
* Click Live to live-view with the camera for making further adjustments
* Raise the light source and exposure times and see the signal becoming higher than the noise, explain why/that it should be higher than the noise
* Then lower the both down and see the signal disappearing in the noise (undesirable in a real experiment)
* Save the light and exposure time adjustments using right-click on the Optical Configuration and choosing "Assign" - explain that this is necessary upon changing something
* Select another pre-set (e.g. mCherry) and point out that another fluorescent color will be activated now and why
* Move on from Epifluorescence-by-eye mode as this is not worth spending the users' brain power on too much.

Confocal Spinning Disc mode (in-depth camera settings)

SoRa Superresolution to double resolving power

ND Acquisition for automated multidimensional experiments, start with Z-stack because full SoRa pipeline needs this.

Perfect Focus System for keeping/using exact focus levels in live view and during ND Acquisition experiments

FRAP (optional)

Dual camera mode (optional)

Data saving (nd2 files) and transfering (SURF file sender)

User account creation

Booking system

Nikon sora introduction protocol

2026-05-27T16:26:45Z

Jelle:

'''Introduction protocol'''

Type of microscope and its strengths

* Spinning disc microscope for fast confocal microscopy (optical sectioning)
* Suitable for live-cell imaging experiments
* Superresolution optional, to achieve (ideally) ca. 150 nm resolution in green.
* Two-camera option for capturing two fluorescent colors at the same time (dynamic processes)
* Fluorescence Recovery After Photobleaching (FRAP) to study molecular diffusion times

[[Nikon sora switch on|Switch-on procedure]]

* Make sure to also start the software already and start Single-cam mode

Stage-top incubation system for live-cell imaging (switch on before starting a live cell experiment)

* Component 2 is the "Warming box" and needs to be switched on as well as the "Heater" button on its front panel
* Look at the stage-top incubator itself - it has a lid that should be used during the experiment
* There is a plastic removable frame covering a "canal" (basin) of water around the edge of the sample area
* The "canal" should be filled with MQ water using the attached tubing - the tubing sticks out of the front of the microscope
* Unclip the tubing clip, attach a syringe with MQ water fill stage-top incubator "canal" with MQ water
* Clip the tubing clip again, remove the syringe, and cover the canal with the plastic frame. It is ready to use

First manual operation

* Operate the lens switcher at the front of the microscope and select the 100X / Oil lens.
* Operate the joystick in various directions as an example. You may have to switch its movement speed to fast.
* Center the stage above the objective.
* Operate the focus knob on the side of the joystick to see the objective move up and down (also may have to switch the movement speed to fast).
* Lower the lens as far as possible down to prepare for placing the sample.

First brightfield operation

* Mount a demo sample (or user sample) in the appropriate sample holder without placing it in the microscope yet
* Put oil on the lens
* Put the condenser arm in the backward position (access to sample area easier)
* Mount the sample holder with sample onto the microscope stage
* Make sure the condenser arm (top arm of the microscope) is set to "O" (brightfield shutter = Open) and not "C"
* Restore the condenser arm to the normal operating position (upright, not backwards)
* Switch on the brightfield lamp (lamp icon, left side of the microscope body) and demonstrate adjusting light intensity (left side rotatable adjuster knob)
* Make sure the eye-piece is active using the "eye" button at the front of the microscope
* Confirm with the user that light is visible through the microscope and that it responds to intensity adjustment
* Visually point the brightfield-lamp-spot onto a relevant sample area, where we expect to see structures
* Let the user raise the objective while looking through the eyepieces, until the sample becomes visible

Epifluorescence mode by camera (quick checking fluorescent signals)

* Go to the computer and make sure everyone can see the screen
* Point out that the software looks complicated but we will discuss the essential parts to understand them
* Roughly describe that there are modalities (Eye-observation, Epifluorescence with camera, Spinning-disc modalities).
* Point out that we were using the Eye-observation mode all this time
* Switch to Epifluorescence with camera mode
* Choose a pre-set optical configuration that should reveal a fluorescent color that is in the sample
* Point out that the two settings that matter now are a light intensity slider (lower left) and a camera exposure time (top left), set them both to low values
* Click Capture image and probably see nothing (black image, but it may just be a low amount of signal!)
* Switch on auto-scale (above the picture, a graph symbol with a Play button on top of it) and explain that this will always show what the camera sees, even if it is weak or strong signal. Now you will see something.
* Click Live to live-view with the camera for making further adjustments
* Raise the light source and exposure times and see the signal becoming higher than the noise, explain why/that it should be higher than the noise
* Then lower the both down and see the signal disappearing in the noise (undesirable in a real experiment)
* Save the light and exposure time adjustments using right-click on the Optical Configuration and choosing "Assign" - explain that this is necessary upon changing something
* Select another pre-set (e.g. mCherry) and point out that another fluorescent color will be activated now and why
* Move on from Epifluorescence-by-eye mode as this is not worth spending the users' brain power on too much.

Confocal Spinning Disc mode (in-depth camera settings)

SoRa Superresolution to double resolving power

ND Acquisition for automated multidimensional experiments, start with Z-stack because full SoRa pipeline needs this.

Perfect Focus System for keeping/using exact focus levels in live view and during ND Acquisition experiments

FRAP (optional)

Dual camera mode (optional)

Data saving (nd2 files) and transfering (SURF file sender)

User account creation

Booking system

Nikon sora introduction protocol

2026-05-27T16:08:00Z

Jelle:

Introduction protocol

Type of microscope and its strengths

* Spinning disc microscope for fast confocal microscopy (optical sectioning)
* Suitable for live-cell imaging experiments
* Superresolution optional, to achieve (ideally) ca. 150 nm resolution in green.
* Two-camera option for capturing two fluorescent colors at the same time (dynamic processes)
* Fluorescence Recovery After Photobleaching (FRAP) to study molecular diffusion times

Stage-top incubation system for live-cell imaging (switch on before starting a live cell experiment)

First manual operation and doing Brightfield (finding the sample)

Epifluorescence mode by eye and by camera (quick checking fluorescent signals)
Confocal Spinning Disc mode (in-depth camera settings)

SoRa Superresolution to double resolving power

ND Acquisition for automated multidimensional experiments, start with Z-stack because full SoRa pipeline needs this.

Perfect Focus System for keeping/using exact focus levels in live view and during ND Acquisition experiments

FRAP (optional)

Dual camera mode (optional)

Data saving (nd2 files) and transfering (SURF file sender)

User account creation

Booking system discussion

Nikon sora introduction protocol

2026-05-27T16:07:41Z

Jelle:

Introduction protocol

Type of microscope and its strengths

* Spinning disc microscopy for fast confocal microscopy (optical sectioning)
* Suitable for live-cell imaging experiments
* Superresolution optional, to achieve (ideally) ca. 150 nm resolution in green.
* Two-camera option for capturing two fluorescent colors at the same time (dynamic processes)
* Fluorescence Recovery After Photobleaching (FRAP) to study molecular diffusion times

Stage-top incubation system for live-cell imaging (switch on before starting a live cell experiment)

First manual operation and doing Brightfield (finding the sample)

Epifluorescence mode by eye and by camera (quick checking fluorescent signals)
Confocal Spinning Disc mode (in-depth camera settings)

SoRa Superresolution to double resolving power

ND Acquisition for automated multidimensional experiments, start with Z-stack because full SoRa pipeline needs this.

Perfect Focus System for keeping/using exact focus levels in live view and during ND Acquisition experiments

FRAP (optional)

Dual camera mode (optional)

Data saving (nd2 files) and transfering (SURF file sender)

User account creation

Booking system discussion

Nikon sora introduction protocol

2026-05-26T15:55:17Z

Jelle:

Introduction protocol

* Type of microscope and its strengths
* Spinning disc microscopy for fast confocal microscopy (optical sectioning)
* Suitable for live-cell imaging experiments
* Superresolution optional, to achieve (ideally) ca. 150 nm resolution in green.
* Two-camera option for capturing two fluorescent colors at the same time (dynamic processes)
* Fluorescence Recovery After Photobleaching (FRAP) to study molecular diffusion times

Stage-top incubation system for live-cell imaging (switch on before starting a live cell experiment)

First manual operation and doing Brightfield (finding the sample)

Epifluorescence mode by eye and by camera (quick checking fluorescent signals)
Confocal Spinning Disc mode (in-depth camera settings)

SoRa Superresolution to double resolving power

ND Acquisition for automated multidimensional experiments, start with Z-stack because full SoRa pipeline needs this.

Perfect Focus System for keeping/using exact focus levels in live view and during ND Acquisition experiments

FRAP (optional)

Dual camera mode (optional)

Data saving (nd2 files) and transfering (SURF file sender)

User account creation

Booking system discussion

Nikon sora introduction protocol

2026-05-26T15:54:15Z

Jelle:

Introduction protocol

Type of microscope and its strengths

- Spinning disc microscopy for fast confocal microscopy (optical sectioning)
- Suitable for live-cell imaging experiments
- Superresolution optional, to achieve (ideally) ca. 150 nm resolution in green.
- Two-camera option for capturing two fluorescent colors at the same time (dynamic processes)
- Fluorescence Recovery After Photobleaching (FRAP) to study molecular diffusion times

Stage-top incubation system for live-cell imaging (switch on before starting a live cell experiment)
First manual operation and doing Brightfield (finding the sample)
Epifluorescence mode by eye and by camera (quick checking fluorescent signals)
Confocal Spinning Disc mode (in-depth camera settings)
SoRa Superresolution to double resolving power
ND Acquisition for automated multidimensional experiments, start with Z-stack because full SoRa pipeline needs this.
Perfect Focus System for keeping/using exact focus levels in live view and during ND Acquisition experiments

FRAP (optional)
Dual camera mode (optional)

Data saving (nd2 files) and transfering (SURF file sender)
User account creation
Booking system discussion

Nikon sora

2026-05-26T15:53:46Z

Jelle: correct introduction protocol link

== Manuals and user guides ==
[[Nikon sora functions|What can it do?]]

[[Nikon sora introduction protocol|Introduction protocol]]

[[Nikon sora switch on|How to switch on]]

[[Nikon sora superresolution switch|How to switch to superresolution]]

[[Nikon sora lightpath|Dual camera imaging wavelengths]]

[[Nikon sora tips tricks|Tips & Tricks]]

== Current status ==
The current status of the microscope can be found [[Current microscope status#Light microscopes|here]] and on the [https://bookings.science.ru.nl/category/general-instrumentation-gi/gi-light-microscopy-image-analysis booking website].

== Biosafety and access ==
The SoRa is located in an ML-1 room. See the [[General Instrumentation Department#Biosafety|Biosafety]] section on rules regarding working in this room, and how to get access.

== Technical specifications ==

=== Available hardware ===

* 20x air obj.
* 40x/1.30 oil (working distance 200 um)
* 100x/1.49 oil (w.d. 160 um)

=== Superresolution components ===

* SoRa spinning disc
* Additional SoRa magnifiers 2.8x / 4.0x
* 3D deconvolution software (in-line and parallel)

=== Imaging lasers ===

* 405 nm
* 445 nm
* 488 nm
* 561 nm
* 638 nm

=== Emission filters ===

* UV
* Green
* Yellow
* Red
* Far-red

=== Dual color simultaneous imaging ===

* blue/yellow (e.g. CFP / YFP)
* green/red (e.g. GFP / RFP)

=== Software ===

* NIS-elements
* Nikon batch-deconvolution

=== Other ===

* FURA (calcium sensor) imaging filters and light source
* "Tokai Hit" Stage-top live-cell incubation system (CO2, temperature and humidity control)

Nikon sora

2026-05-26T15:52:58Z

Jelle: added introduction protocol link

== Manuals and user guides ==
[[Nikon sora functions|What can it do?]]

[[Introduction protocol]]

[[Nikon sora switch on|How to '''switch on''']]

[[Nikon sora superresolution switch|How to '''switch to superresolution''']]

[[Nikon sora lightpath|Dual camera imaging wavelengths]]

[[Nikon sora tips tricks|Tips & Tricks]]

== Current status ==
The current status of the microscope can be found [[Current microscope status#Light microscopes|here]] and on the [https://bookings.science.ru.nl/category/general-instrumentation-gi/gi-light-microscopy-image-analysis booking website].

== Biosafety and access ==
The SoRa is located in an ML-1 room. See the [[General Instrumentation Department#Biosafety|Biosafety]] section on rules regarding working in this room, and how to get access.

== Technical specifications ==

=== Available hardware ===

* 20x air obj.
* 40x/1.30 oil (working distance 200 um)
* 100x/1.49 oil (w.d. 160 um)

=== Superresolution components ===

* SoRa spinning disc
* Additional SoRa magnifiers 2.8x / 4.0x
* 3D deconvolution software (in-line and parallel)

=== Imaging lasers ===

* 405 nm
* 445 nm
* 488 nm
* 561 nm
* 638 nm

=== Emission filters ===

* UV
* Green
* Yellow
* Red
* Far-red

=== Dual color simultaneous imaging ===

* blue/yellow (e.g. CFP / YFP)
* green/red (e.g. GFP / RFP)

=== Software ===

* NIS-elements
* Nikon batch-deconvolution

=== Other ===

* FURA (calcium sensor) imaging filters and light source
* "Tokai Hit" Stage-top live-cell incubation system (CO2, temperature and humidity control)

Nikon sora introduction protocol

2026-05-26T15:52:04Z

Jelle: created page

Introduction protocol

Type of microscope and its strengths
- Spinning disc microscopy for fast confocal microscopy (optical sectioning)
- Suitable for live-cell imaging experiments
- Superresolution optional, to achieve (ideally) ca. 150 nm resolution in green.
- Two-camera option for capturing two fluorescent colors at the same time (dynamic processes)
- Fluorescence Recovery After Photobleaching (FRAP) to study molecular diffusion times

Stage-top incubation system for live-cell imaging (switch on before starting a live cell experiment)
First manual operation and doing Brightfield (finding the sample)
Epifluorescence mode by eye and by camera (quick checking fluorescent signals)
Confocal Spinning Disc mode (in-depth camera settings)
SoRa Superresolution to double resolving power
ND Acquisition for automated multidimensional experiments, start with Z-stack because full SoRa pipeline needs this.
Perfect Focus System for keeping/using exact focus levels in live view and during ND Acquisition experiments

FRAP (optional)
Dual camera mode (optional)

Data saving (nd2 files) and transfering (SURF file sender)
User account creation
Booking system discussion

Nikon TIRF widefield

2025-11-24T15:17:28Z

Jelle: /* Technical specifications */

== Manuals and user guides ==
[[TIRF alignment guide]]
== Current status ==
The current status of the microscope can be found [[Current microscope status#Light microscopes|here]] and on the [https://bookings.science.ru.nl/category/general-instrumentation-gi/gi-light-microscopy-image-analysis booking website].
== Biosafety and access ==
The TIRF is located in an ML-1 room. See the [[General Instrumentation Department#Biosafety|Biosafety]] section on rules regarding working in this room, and how to get access.

== Technical specifications ==
'''Light sources and filters:'''

'''Epifluorescence mode''' with LED light sources (''non-TIRF)'': Excitations: 385, 475, 550, 621 nm. Emissions: Band pass emission filters for blue (ca. 450), green (ca. 510), red (ca. 600) and far-red (ca. 670).

'''TIRF-mode''' with lasers: Excitations: 488 nm, 561 nm, 640 nm. Emission filters: None

Current microscope status

2025-11-20T15:14:13Z

Jelle:

==== Department closure during the Christmas holiday (update 14.11) ====

* The Huygens building and therefor the GI department will be closed during the Christmas holiday from Monday 22 December up to and including January 2nd.
** Only people with special card access can enter the GI department. Remember to use the overwork timer!
** No support staff will be on site.
** Access to liquid nitrogen is not guaranteed.
* On January 5-6-7 the Huygens building will inaccessible due to maintenance work. No one will be allowed inside; this includes people with extra card access.

=== Light microscopes ===
{| class="wikitable"
|+
!Microscope
!Status
!Notes
!Date
|-
|[[Leica SP8X|SP8X]]
|OK
|
|
|-
|[[Leica SP8 Liachroic|SP8-Lia]]
|OK
|488 laser functional again
|20.11.2025
|-
|[[Leica Thunder|Thunder]]
|OK
|
|
|-
|[[Leica Timelapse|Timelapse]]
|OK
|New camera installed
|25.09.2025
|-
|[[PhaseView light sheet|Lightsheet]]
|
|Microscope fully functional, use 4TB SSD drive to take images
|20.11.2025
|-
|[[DM2500]]
|OK
|
|
|-
|[[Nikon sora|SoRa]]
|OK
|
|
|-
|[[Nikon TIRF widefield|TIRF]]
|OK
|
|
|-
|Spinoza server
|OK
|
|
|}

=== Electron microscopes and sample prep equipment ===
{| class="wikitable"
|+
!Machine
!Status
!Notes
!Date
|-
|[[JEOL1400 TEM|JEOL1400]]
|OK
|
|
|-
|[[JEOL2100 TEM|JEOL2100]]
|Broken
|HT problems. Contacted JEOL.
No date as of yet, but will not happen in November

|12.11.2025
14.11.2025
|-
|
|
|STEM detector broken - under repair
EDX detector broken
|aug 2024
pre 2023
|-
|[[Cryo-holder, cryo-dock and cryo-pumping station|Cryo-dock/station/holder]]
|OK
|
|
|-
|[[HPM100]]
|OK
|
|
|-
|[[EMPACT|EMpact]]
|Broken
|Unsure if can be fixed
|01.09.2025
|-
|[[Vitrobot]]
|OK
|
|
|-
|[[UC-E microtome]]
|OK
|
|
|-
|UC7/FC7 cryo-microtome
|OK
|
|
|-
|[[CCU-010]]
|OK
|
|
|-
|Edward306
|Broken
|Under repair
|
|-
|BAF400
|Broken
|Under repair
|
|-
|Vibratome
|OK
|
|
|-
|CPD-020
|In storage
|
|
|-
|JFD-030
|OK
|
|
|}

Nikon sora tips tricks

2025-10-16T07:51:58Z

Jelle:

'''Tips & Tricks'''

- The function "'''Auto-Capture'''" captures an image and autosaves it to the specified folder (configured in Auto-Capture Settings).

-

Nikon sora

2025-10-16T07:51:40Z

Jelle:

== Manuals and user guides ==
[[Nikon sora functions|What can it do?]]

[[Nikon sora switch on|How to '''switch on''']]

[[Nikon sora superresolution switch|How to '''switch to superresolution''']]

[[Nikon sora lightpath|Dual camera imaging wavelengths]]

[[Nikon sora tips tricks|Tips & Tricks]]

== Current status ==
The current status of the microscope can be found [[Current microscope status#Light microscopes|here]] and on the [https://bookings.science.ru.nl/category/general-instrumentation-gi/gi-light-microscopy-image-analysis booking website].

== Biosafety and access ==
The SoRa is located in an ML-1 room. See the [[General Instrumentation Department#Biosafety|Biosafety]] section on rules regarding working in this room, and how to get access.

== Technical specifications ==

=== Available hardware ===

* 20x air obj.
* 40x/1.30 oil (working distance 200 um)
* 100x/1.49 oil (w.d. 160 um)

=== Superresolution components ===

* SoRa spinning disc
* Additional SoRa magnifiers 2.8x / 4.0x
* 3D deconvolution software (in-line and parallel)

=== Imaging lasers ===

* 405 nm
* 445 nm
* 488 nm
* 561 nm
* 638 nm

=== Emission filters ===

* UV
* Green
* Yellow
* Red
* Far-red

=== Dual color simultaneous imaging ===

* blue/yellow (e.g. CFP / YFP)
* green/red (e.g. GFP / RFP)

=== Software ===

* NIS-elements
* Nikon batch-deconvolution

=== Other ===

* FURA (calcium sensor) imaging filters and light source
* "Tokai Hit" Stage-top live-cell incubation system (CO2, temperature and humidity control)

Nikon sora tips tricks

2025-10-16T07:51:06Z

Jelle: Created page with "Tips & Tricks - The function "'''Auto-Capture'''" captures an image and autosaves it to the specified folder (configured in Auto-Capture Settings). -"

Tips & Tricks

- The function "'''Auto-Capture'''" captures an image and autosaves it to the specified folder (configured in Auto-Capture Settings).

-

Current microscope status

2025-10-03T11:14:27Z

Jelle:

=== Light microscopes ===
{| class="wikitable"
|+
!Microscope
!Status
!Notes
!Date
|-
|[[Leica SP8X|SP8X]]
|OK
|
|
|-
|[[Leica SP8 Liachroic|SP8-Lia]]
|OK
|488 laser dodgy - don't turn it off!
|27.08.2025
|-
|[[Leica Thunder|Thunder]]
|OK
|
|
|-
|[[Leica Timelapse|Timelapse]]
|OK
|New camera installed
|25.09.2025
|-
|[[PhaseView light sheet|Lightsheet]]
|
|561 laser weak+ software out of date
Broken parts are sent for repair (PhaseView corp., Paris, France)
|03.10.2025
|-
|[[DM2500]]
|OK
|
|
|-
|[[Nikon sora|SoRa]]
|OK
|
|
|-
|[[Nikon TIRF widefield|TIRF]]
|OK
|
|
|-
|Spinoza server
|OK
|
|
|}

=== Electron microscopes and sample prep equipment ===
{| class="wikitable"
|+
!Machine
!Status
!Notes
!Date
|-
|[[JEOL1400 TEM|JEOL1400]]
|OK
|
|
|-
|[[JEOL2100 TEM|JEOL2100]]
|OK
|STEM detector broken - under repair
EDX detector broken
|aug 2024
pre 2023
|-
|[[Cryo-holder, cryo-dock and cryo-pumping station|Cryo-dock/station/holder]]
|OK
|
|
|-
|[[HPM100]]
|OK
|
|
|-
|[[EMPACT|EMpact]]
|Broken
|Unsure if can be fixed
|01.09.2025
|-
|[[Vitrobot]]
|OK
|
|
|-
|[[UC-E microtome]]
|Broken
|Under repair - contact Hetty if you need it soon
|01.09.2025
|-
|UC7/FC7 cryo-microtome
|OK
|
|
|-
|[[CCU-010]]
|OK
|
|
|-
|Edward306
|Broken
|Under repair
|
|-
|BAF400
|Broken
|Under repair
|
|-
|Vibratome
|OK
|
|
|-
|
|
|
|
|}

Current microscope status

2025-10-03T11:14:14Z

Jelle:

=== Light microscopes ===
{| class="wikitable"
|+
!Microscope
!Status
!Notes
!Date
|-
|[[Leica SP8X|SP8X]]
|OK
|
|
|-
|[[Leica SP8 Liachroic|SP8-Lia]]
|OK
|488 laser dodgy - don't turn it off!
|27.08.2025
|-
|[[Leica Thunder|Thunder]]
|OK
|
|
|-
|[[Leica Timelapse|Timelapse]]
|OK
|New camera installed
|25.09.2025
|-
|[[PhaseView light sheet|Lightsheet]]
|
|561 laser weak+ software out of date
Broken parts are sent for repair (PhaseView corp., Paris, France)
|?
|-
|[[DM2500]]
|OK
|
|
|-
|[[Nikon sora|SoRa]]
|OK
|
|
|-
|[[Nikon TIRF widefield|TIRF]]
|OK
|
|
|-
|Spinoza server
|OK
|
|
|}

=== Electron microscopes and sample prep equipment ===
{| class="wikitable"
|+
!Machine
!Status
!Notes
!Date
|-
|[[JEOL1400 TEM|JEOL1400]]
|OK
|
|
|-
|[[JEOL2100 TEM|JEOL2100]]
|OK
|STEM detector broken - under repair
EDX detector broken
|aug 2024
pre 2023
|-
|[[Cryo-holder, cryo-dock and cryo-pumping station|Cryo-dock/station/holder]]
|OK
|
|
|-
|[[HPM100]]
|OK
|
|
|-
|[[EMPACT|EMpact]]
|Broken
|Unsure if can be fixed
|01.09.2025
|-
|[[Vitrobot]]
|OK
|
|
|-
|[[UC-E microtome]]
|Broken
|Under repair - contact Hetty if you need it soon
|01.09.2025
|-
|UC7/FC7 cryo-microtome
|OK
|
|
|-
|[[CCU-010]]
|OK
|
|
|-
|Edward306
|Broken
|Under repair
|
|-
|BAF400
|Broken
|Under repair
|
|-
|Vibratome
|OK
|
|
|-
|
|
|
|
|}

Current microscope status

2025-10-03T11:13:23Z

Jelle:

=== Light microscopes ===
{| class="wikitable"
|+
!Microscope
!Status
!Notes
!Date
|-
|[[Leica SP8X|SP8X]]
|OK
|
|
|-
|[[Leica SP8 Liachroic|SP8-Lia]]
|OK
|488 laser dodgy - don't turn it off!
|27.08.2025
|-
|[[Leica Thunder|Thunder]]
|OK
|
|
|-
|[[Leica Timelapse|Timelapse]]
|OK
|New camera installed
|25.09.2025
|-
|[[PhaseView light sheet|Lightsheet]]
|
|561 laser weak+ software out of date
Broken microscope parts are sent for repair (PhaseView corp., Paris, France)
|?
|-
|[[DM2500]]
|OK
|
|
|-
|[[Nikon sora|SoRa]]
|OK
|
|
|-
|[[Nikon TIRF widefield|TIRF]]
|OK
|
|
|-
|Spinoza server
|OK
|
|
|}

=== Electron microscopes and sample prep equipment ===
{| class="wikitable"
|+
!Machine
!Status
!Notes
!Date
|-
|[[JEOL1400 TEM|JEOL1400]]
|OK
|
|
|-
|[[JEOL2100 TEM|JEOL2100]]
|OK
|STEM detector broken - under repair
EDX detector broken
|aug 2024
pre 2023
|-
|[[Cryo-holder, cryo-dock and cryo-pumping station|Cryo-dock/station/holder]]
|OK
|
|
|-
|[[HPM100]]
|OK
|
|
|-
|[[EMPACT|EMpact]]
|Broken
|Unsure if can be fixed
|01.09.2025
|-
|[[Vitrobot]]
|OK
|
|
|-
|[[UC-E microtome]]
|Broken
|Under repair - contact Hetty if you need it soon
|01.09.2025
|-
|UC7/FC7 cryo-microtome
|OK
|
|
|-
|[[CCU-010]]
|OK
|
|
|-
|Edward306
|Broken
|Under repair
|
|-
|BAF400
|Broken
|Under repair
|
|-
|Vibratome
|OK
|
|
|-
|
|
|
|
|}

Leica SP8X

2025-09-30T10:55:05Z

Jelle:

The SP8x confocal laser scanning microscope (Leica-microsystems) is mounted on a fully motorized DMi8 microscope on which x, y, z, t; lambda (wavelength), and sequential scans can be acquired at multiple positions and in z-stack. There are functions for stitching, and digital “best focus” determination also in z-stacks. Deconvolution can be done on the attached CUDA PC equipped with the LAS X module "Lightning". Live-cell imaging can be performed using a closed lid that controls CO2 and relative humidity levels, and using the big incubator box that controls temperature and airflow.

== Manuals and user guides ==

== Current status ==
The current status of the microscope can be found [[Current microscope status#Light microscopes|here]] and on the [https://bookings.science.ru.nl/category/general-instrumentation-gi/gi-light-microscopy-image-analysis booking website].

== Biosafety and access ==
The SP8-Lia is located in an AP-1 room. See the [[General Instrumentation Department#Biosafety|Biosafety]] section on rules regarding working in this room.

== Technical specifications ==

=== Objectives ===

* 10x / 0.40 Air (FWD = 2.2 mm)
* 40x / 0.60 Air (3.3 mm long distance)
* 40x / 1.10 Water (0.62 mm) - motorCorrection
* 63x / 1.40 Oil (0.14 mm)
* 100x / 1.40 Oil (0.13 mm)

===Lasers===
* 405 nm diode laser + pulsed White Light Laser (WLL), up to 8 tunable excitation lines from 470-670nm can be applied simultaneously

===Detectors and camera===
* 2 GaAsP PMTs
* 2 HyDs (high sensitivity, linear photon counting and time-gating)
* 1 normal transmitted-light PMT

In addition, the DMi8 microscope can be utilized for wide-field bright field and fluorescence observation. Images can be acquired with a monochrome DFC365FX camera.

===Filter cubes===
* DAPI (LP425)
* FITC (LP515)
* Rhod (LP590)
* Y5 (BP 662-738)

Current microscope status

2025-09-26T09:20:42Z

Jelle:

=== Light microscopes ===
{| class="wikitable"
|+
!Microscope
!Status
!Notes
!Date
|-
|SP8X
|OK
|
|
|-
|[[Leica SP8 Liachroic|SP8-Lia]]
|OK
|488 laser dodgy - don't turn it off!
|27.08.2025
|-
|Thunder
|OK
|
|
|-
|Timelapse
|OK
|New camera installed
|25.09.2025
|-
|Lightsheet
|
|561 laser weak
Software out of date

service visit planned 03.10.2025
|?
|-
|DM2500
|OK
|
|
|-
|[[Nikon sora|SoRa]]
|OK
|
|
|-
|[[Nikon TIRF widefield|TIRF]]
|OK
|
|
|-
|Spinoza server
|OK
|
|
|}

=== Electron microscopes and sample prep equipment ===
{| class="wikitable"
|+
!Machine
!Status
!Notes
!Date
|-
|[[JEOL1400 TEM|JEOL1400]]
|OK
|
|
|-
|JEOL2100
|OK
|STEM detector broken - under repair
EDX detector broken
|aug 2024
pre 2023
|-
|Cryo-dock/station/holder
|OK
|
|
|-
|[[HPM100]]
|OK
|
|
|-
|[[EMPACT|EMpact]]
|Broken
|Unsure if can be fixed
|01.09.2025
|-
|Vitrobot
|OK
|
|
|-
|UC-E
|Broken
|Under repair - contact Hetty if you need it soon
|01.09.2025
|-
|UC7/FC7 cryo-microtome
|OK
|
|
|-
|[[CCU-010]]
|OK
|
|
|-
|Edward306
|Broken
|Under repair
|
|-
|BAF400
|Broken
|Under repair
|
|-
|Vibratome
|OK
|
|
|-
|
|
|
|
|}

Current microscope status

2025-09-26T09:20:32Z

Jelle:

=== Light microscopes ===
{| class="wikitable"
|+
!Microscope
!Status
!Notes
!Date
|-
|SP8X
|OK
|
|
|-
|[[Leica SP8 Liachroic|SP8-Lia]]
|OK
|488 laser dodgy - don't turn it off!
|27.08.2025
|-
|Thunder
|OK
|
|
|-
|Timelapse
|OK
|New camera installed
|25.09.2025
|-
|Lightsheet
|
|561 laser weak
Software out of date
service visit planned 03.10.2025
|?
|-
|DM2500
|OK
|
|
|-
|[[Nikon sora|SoRa]]
|OK
|
|
|-
|[[Nikon TIRF widefield|TIRF]]
|OK
|
|
|-
|Spinoza server
|OK
|
|
|}

=== Electron microscopes and sample prep equipment ===
{| class="wikitable"
|+
!Machine
!Status
!Notes
!Date
|-
|[[JEOL1400 TEM|JEOL1400]]
|OK
|
|
|-
|JEOL2100
|OK
|STEM detector broken - under repair
EDX detector broken
|aug 2024
pre 2023
|-
|Cryo-dock/station/holder
|OK
|
|
|-
|[[HPM100]]
|OK
|
|
|-
|[[EMPACT|EMpact]]
|Broken
|Unsure if can be fixed
|01.09.2025
|-
|Vitrobot
|OK
|
|
|-
|UC-E
|Broken
|Under repair - contact Hetty if you need it soon
|01.09.2025
|-
|UC7/FC7 cryo-microtome
|OK
|
|
|-
|[[CCU-010]]
|OK
|
|
|-
|Edward306
|Broken
|Under repair
|
|-
|BAF400
|Broken
|Under repair
|
|-
|Vibratome
|OK
|
|
|-
|
|
|
|
|}

Leica SP8 Liachroic intro

2025-09-23T10:02:50Z

Jelle:

'''Introduction'''

# Welcome, outline of what the introduction will be (takes a few hours)
# The General Instrumentation department where we are
# The room and the general layout (sink, microscope, cupboard, chairs)
# The microscope conceptually (Confocal Laser Scanning Microscope)
## '''Confocal''' = observing only a small horizontal layer of the sample, not seeing blurry higher or lower objects.
## '''Laser''' = specific wavelengths, focused spot, to only see specifically what we want
## '''Scanning''' = doesn't take a picture in one shot, but scans every pixel by pixel until the whole picture of a square field of view is built.
# The microscope physically (components, roughly what parts do we see? -> microscope, scanner (left, attached to microscope on air table), laser box (left next to air table))

'''Switch-on'''

# Quick inspection (everything looks ok visually?)
# Switch-on procedure of components (buttons in numbered order)
# Log-in in Windows (use yours, user will get an account) and software starting (LAS X)

'''Stage and objective movement controls''' (XY stage joystick, Z objectives, Switch Objectives on the touch screen - start with the 10X lens. You normally start low, then go higher)

'''Sample placement''' (sample types, coverslip towards the bottom, adjusting holder, put demo or user sample there, don't clamp it but let it rest (otherwise breaks when objective hits))

'''Brightfield''' (where does light come from, what do we expect to see)

'''Focusing''' (sample finding, and then moving around, warning: hitting lens to sample possible)

'''Epifluorescence''' (see visually light shining, see returning light, discuss different filters, user responsible for knowing their dyes and wavelengths, Searchlight spectra viewer. This method is used to see where is the signal, is there any signal, do I already see something suitable for confocal imaging?)

'''Confocal scanning''' (gonna be a big part, mostly on computer now, switch in mood/user to focus on screen not on microscope - instructor sits on the far-right side of the desk to control the mouse, attendees sit right in front of the screen to see everything properly)

- Launch the software LAS X which controls the microscope, and choose the right startup settings (microscope = DMi8, settings = standard) and wait for it to appear completely. The software is going to ignore any wavelength/imaging modality (BF, EPI) choices you have made using the touch screen of the microscope and do confocal anyway.

'''- Overview of software areas:'''

# We start in "Acquire" (top) and will be spending most time there
# The big vertical line in the middle divides the view into settings (left) and image display (right)
# Making any adjustments to settings in the image display (right) is not going to change any raw data values or any microscope settings - just how the image looks on the screen
# Making adjustments on the left will change actual microscope settings, such as wavelengths used, scanning behavior etc.
# At the bottom, there are the buttons that make the microscope actually do something (Live, Capture Image, Start - etc.) - to prevent confusion when clicking "open laser shutter" in the software, or any other buttons.
# We make 1 small excursion to "Configuration" (left of "Acquire") and then "Laser Config" in order to switch on the lasers we know we will need.

'''- Laser setup'''

# choose the right wavelength
# give it a tiny bit of power (philosophy: start low, go higher if needed - instead of bleaching everything straight away)
# click round button under the laser to open the shutter
# Confirm yellow line appears in the schematic light path below -> laser ready.

'''- Detector setup'''

# Acknowledge we have a few detectors available, one is switched on.
# Prefer to always choose HyD (hybrid detectors), they are better. Typically they are in standard mode with 100 gain (digital gain) and you can use them like that. You can destroy the detector by setting its detection window to include the current laser wavelength used (dotted line in detector area spectrum reminds you which lasers are used).
# PMT detectors are less sensitive = good for strong signals (usually you have weak signals), to use one you have to give it some gain (couple hundred Volts, until image looks good to you)
# Move the detection window and resize it to capture the desired fluorescence emission wavelengths. Avoid including the laser wavelength at all cost!! This will visualise the laser, which we do not want to see - and it can damage the detector.

'''- First image'''

# Click "Capture Image"
# Probably, nothing will show up!
# Is there something there, but just weak? -> Set the Image Display (right) to "auto-scale" (button called "M" because it starts on Manual scale) -> have a look if something shows up.
# Is there nothing there anyway, maybe you need more laser -> Increase laser power to 20-30%, capture another image -> have a look if something shows up.
# Is there still nothing, maybe the focus is slightly wrong (we're in confocal mode after all...) -> use the Z-adjustment knob to focus up and down (either on the microscope, on the joystick, or on the USB panel) -> have a look if something shows up, now it really should.
# If there's still nothing, visually check if the sample is still alright, and then use Brightfield or Epifluorescence through the binoculars to find focus again. Then try confocal again, using the software.
# If there's still nothing, there is a problem with the microscope or the sample. Change samples to a known working sample - a piece of paper responds to 405 nm, a stroke of Staedler Yellow Highlighter responds to 488 nm.

'''- Evaluating the first image, boundary conditions'''

# It could be too weak (barely jumping out of the noise), so we need more laser until it's visible (improving the scan will help too, but this comes later) at the cost of possibly bleaching
# It could be too strong (huge areas look intense, and same intensity - mouse-over the pixel and see the maximum value 255 show up under the picture = too bright!) you want to avoid saturation.

'''- Optimizing the scan (better picture, takes longer, make it as good as you need - and not better. Every improvement you make can be compared to the previous situation by flipping through the last couple of acquired images)'''

# This part is about losing acquisition speed but gaining image quality.
# There are three main settings that most influence the image quality, which you can use to maximize the details and signals that the current objective is able to see. (if not enough, you need a higher mag. objective) as follows:
# '''Format''' (number of pixels that are scanned -> note that the square area of the sample that you are scanning does not change in size, just gets divided into smaller steps i.e. more measurement points i.e. pixels). This influences the resolution of the image (lower = worse). Use the "Optimize" button on the left of Format to always get the right number of pixels to describe the details that are available through this objective.
# '''Speed''' (lower values = slower, takes longer but results in higher signal-to-noise). This means the laser spends more time on each pixel, and thus illuminates that place for longer, and thus gets more signals if there are signals. Noise always stays the same.
# '''Averaging''' (higher values = longer, also more signal-to-noise). This means one horizontal line will be acquired that number of times and the values averaged. Noise randomly appears everywhere. Signal always appears, on every scan consistently, when you look at a location that has signal.

'''- Adding a second channel and using Sequential Scanning'''

# Set the settings to "bad" again (512x512 pixels, 400 speed, no averaging) to save some time.
# Set up another laser (e.g. 561, if you used 488) to activate the other fluorophores
# Set up another detector (e.g. HyD3 if you used HyD1) to detect the other fluorophores' signals
# Do another "Capture Image" and make optimizations where necessary (use the Evaluating the second image just like in the first image section above).
# '''Sequential Scanning''': both molecule populations and both detectors are active at the same time, so bleedthrough might happen.
# Switch on "SEQ" at the top left, a sequential scanning menu will appear.
# Add another sequence using "+" and switch off one of the detectors in sequence 2
# Flip back and forth between seq 1 and 2 so that you can see the difference in settings.
# Now set up sequence 1 for one imaging channel, and sequence 2 for the other: Lasers should be "switched off" by putting their power on 0 where necessary, in each sequence.

'''- Switching to a higher magnification lens (use navigator to register 1 pic, switch to immersion lens, go back to original location).'''

# Higher magnification does not automatically mean higher detail on a confocal scanning microscope.
# Higher NA does mean higher detail on a confocal. Provided that the scan settings are also optimized (just done that, can repeat it).

'''- Z-stack''' (look at the menu: set start height, go to a different focus and set end height, and that's it - would always recommend to leave the rest of the Z-stack settings as-is for the introduction)

'''- Time course''' (switch modes using dropdown menu, then set (1) interval (none, or amount of time) and (2) duration (unlimited, or amount of time, or amount of pictures)

'''- Tilescan''' (Use the Navigator for this, do one small one, several of them, focus map, stitching)

# Make sure some objects are in the view in normal viewer
# Switch to '''Navigator''' and do "Spiral"
# After an area is revealed click "'''Stop'''"
# Use a '''drawing tool''' (bottom, e.g. the rectangle) to designate a small tilescan area
# '''Execute''' ("Start") the tilescan
# In the left menu "Stage", confirm that "Merge images during acquisition" is on, and "blend" is on "Smooth"
# Inspect the result, be skeptical of overlap areas, they should be stitched well

'''Rounding off'''

- Saving data (save the projects as LIF files locally, then before ending your session move the raw data to a safe location e.g. through SURF or network drive, our policy is that the computer is not a safe storage space and we can't count on it)

- Saving settings (sequential scan settings in the sequential scan menu, single settings above the laser set-up area, or you can reload settings from an image in a LIF file: right-click on one and "apply".

- Closing software (is someone coming after us? do we need to keep the whole system on just logging out, or switch the whole system off?)

- Cleaning oil objective, then put back at 10X

- Shutting down microscope (if necessary, otherwise just logging out)

- Leaving room and system tidy for next user!

'''Optional functions'''

- Autofocus

- Emission wavelength scanning (check whether the signal behaves like the fluorophore should)

- Closing the pinhole (more resolution, lower signal)

- Changing Z-stack settings like step size (speed thigns up, but you may skip layers and thus lose information)

Nikon sora lightpath

2025-09-05T09:57:43Z

Jelle:

== Dual camera imaging situation and choices: ==
[[File:Dual_cam_light_path.png|836x836px]]

Nikon sora

2025-09-05T09:57:33Z

Jelle:

[[Nikon sora functions|What can it do?]]

[[Nikon sora switch on|How to '''switch on''']]

[[Nikon sora superresolution switch|How to '''switch to superresolution''']]

[[Nikon sora lightpath|Dual camera imaging wavelengths]]

Nikon sora

2025-09-05T09:57:25Z

Jelle: added dual cam lightpath

Nikon sora lightpath

2025-09-05T09:55:53Z

Jelle:

'''Dual camera imaging situation and choices:'''

[[File:Dual_cam_light_path.png|836x836px]]

Nikon sora lightpath

2025-09-05T09:53:47Z

Jelle: Created page with "thumb"

[[File:Dual cam light path.png|thumb]]

File:Dual cam light path.png

2025-09-05T09:53:42Z

Jelle:

Nikon sora dual camera light path choices

Leica SP8 Liachroic intro

2025-08-14T12:20:54Z

Jelle: additions

'''Introduction'''

# Welcome, outline of the introduction
# The General Instrumentation department conceptually and physically
# The room and the general layout
# The microscope conceptually (functions)
# The microscope physically (components, roughly)

'''Switch-on'''

# Quick inspection (everything looks ok visually?)
# Switch-on procedure of components (buttons numbered order)
# Log-in in Windows (user will get an account) and software starting

'''Stage and objective movement controls''' (XY stage, Z objectives, Switch Objectives on the touch screen - start with the 10X lens. You normally start low, then go higher)

'''Sample placement''' (sample types, coverslip towards the bottom, adjusting holder, put demo or user sample there, don't clamp it but let it rest (otherwise breaks when objective hits))

'''Brightfield''' (where does light come from, what do we expect to see)

'''Focusing''' (sample finding, and then moving around, warning: hitting lens to sample possible)

'''Epifluorescence''' (see visually light shining, see returning light, discuss different filters, user responsible for knowing their dyes and wavelengths, Searchlight spectra viewer. This method is used to see where is the signal, is there any signal, do I already see something suitable for confocal imaging?)

'''Confocal scanning''' (gonna be a big part, mostly on computer now, switch in mood/user to focus on screen not on microscope - instructor sits on the far-right side of the desk to control the mouse, attendees sit right in front of the screen to see everything properly)

- Launch the software LAS X which controls the microscope, and choose the right startup settings (microscope = DMi8, settings = standard) and wait for it to appear completely. The software is going to ignore any wavelength/imaging modality (BF, EPI) choices you have made using the touch screen of the microscope and do confocal anyway.

- Overview of software areas:

# We start in "Acquire" (top) and will be spending most time there
# The big vertical line in the middle divides the view into settings (left) and image display (right)
# Making any adjustments to settings in the image display (right) is not going to change any raw data values or any microscope settings - just how the image looks on the screen
# Making adjustments on the left will change actual microscope settings, such as wavelengths used, scanning behavior etc.
# At the bottom, there are the buttons that make the microscope actually do something (Live, Capture Image, Start - etc.) - to prevent confusion when clicking "open laser shutter" in the software, or any other buttons.
# We make 1 small excursion to "Configuration" (left of "Acquire") and then "Laser Config" in order to switch on the lasers we know we will need.

- Laser setup

# choose the right wavelength
# give it a tiny bit of power (philosophy: start low, go higher if needed - instead of bleaching everything straight away)
# click round button under the laser to open the shutter
# Confirm yellow line appears in the schematic light path below -> laser ready.

- Detector setup

# Acknowledge we have a few detectors available, one is switched on.
# Prefer to always choose HyD (hybrid detectors), they are better. Typically they are in standard mode with 100 gain (digital gain) and you can use them like that. You can destroy the detector by setting its detection window to include the current laser wavelength used (dotted line in detector area spectrum reminds you which lasers are used).
# PMT detectors are less sensitive = good for strong signals (usually you have weak signals), to use one you have to give it some gain (couple hundred Volts, until image looks good to you)
# Move the detection window and resize it to capture the desired fluorescence emission wavelengths. Avoid including the laser wavelength at all cost!! This will visualise the laser, which we do not want to see - and it can damage the detector.

- First image

# Click "Capture Image"
# Probably, nothing will show up!
# Is there something there, but just weak? -> Set the Image Display (right) to "auto-scale" (button called "M" because it starts on Manual scale) -> have a look if something shows up.
# Is there nothing there anyway, maybe you need more laser -> Increase laser power to 20-30%, capture another image -> have a look if something shows up.
# Is there still nothing, maybe the focus is slightly wrong (we're in confocal mode after all...) -> use the Z-adjustment knob to focus up and down (either on the microscope, on the joystick, or on the USB panel) -> have a look if something shows up, now it really should.
# If there's still nothing, visually check if the sample is still alright, and then use Brightfield or Epifluorescence through the binoculars to find focus again. Then try confocal again, using the software.
# If there's still nothing, there is a problem with the microscope or the sample. Change samples to a known working sample - a piece of paper responds to 405 nm, a stroke of Staedler Yellow Highlighter responds to 488 nm.

- Evaluating the first image, boundary conditions

# It could be too weak (barely jumping out of the noise), so we need more laser until it's visible (improving the scan will help too, but this comes later) at the cost of possibly bleaching
# It could be too strong (huge areas look intense, and same intensity - mouse-over the pixel and see the maximum value 255 show up under the picture = too bright!) you want to avoid saturation.

- Optimizing the scan (better picture, takes longer, make it as good as you need - and not better. Every improvement you make can be compared to the previous situation by flipping through the last couple of acquired images)

# There are three main settings that most influence the image quality, which you can use to maximize the details and signals that the current objective is able to see. (if not enough, you need a higher mag. objective)
# Format (number of pixels that are scanned -> note that the square area of the sample that you are scanning does not change in size, just gets divided into smaller steps i.e. more measurement points i.e. pixels). This influences the resolution of the image (lower = worse). Use the "Optimize" button on the left of Format to always get the right number of pixels to describe the details that are available through this objective.
# Speed (lower values = slower, takes longer but results in higher signal-to-noise). This means the laser spends more time on each pixel, and thus illuminates that place for longer, and thus gets more signals if there are signals. Noise always stays the same.

- Adding a second channel

# Set up another laser (e.g. 561, if you used 488) to activate the other fluorophores
# Set up another detector (e.g. HyD3 if you used HyD1) to detect the other fluorophores' signals
# Do another "Capture Image" and make optimizations where necessary (use the Evaluating the first image section above)

- Switching to a higher magnification lens (use navigator to register 1 pic, switch to immersion lens, go back to original location).

# Higher magnification does not automatically mean higher detail on a confocal scanning microscope.
# Higher NA does mean higher detail on a confocal. Provided that the scan settings are also optimized (just done that, can repeat it).

- Z-stack (set start, end, use optimized)

- Time course (switch modes using dropdown menu, then set (1) interval (none, or amount of time) and (2) duration (unlimited, or amount of time, or amount of pictures)

- Tilescan (one small one, several of them, focus map, stitching)

'''Rounding off'''

- Saving data

- Saving settings

- Closing software

- Shutting down microscope (and cleaning oil objective, then put back at 10X)

- Leaving room and system tidy for next user

Optional functions

- Autofocus

- Emission wavelength scanning (check whether the signal behaves like the fluorophore should)

- Closing the pinhole (more resolution, lower signal)

Leica SP8 Liachroic

2025-08-06T16:36:47Z

Jelle: added link

[[Leica SP8 Liachroic intro|Introduction protocol]]

Leica SP8 Liachroic intro

2025-08-06T16:35:53Z

Jelle: drafted most segments and started adding details

'''Introduction'''

# Welcome, outline of the introduction
# The General Instrumentation department conceptually and physically
# The room and the general layout
# The microscope conceptually (functions)
# The microscope physically (components, roughly)

'''Switch-on'''

# Quick inspection (everything looks ok visually?)
# Switch-on procedure of components (buttons numbered order)
# Log-in in Windows (user will get an account) and software starting

'''Stage and objective movement controls''' (XY stage, Z objectives, Switch Objectives on the touch screen - start with the 10X lens. You normally start low, then go higher)

'''Sample placement''' (sample types, coverslip towards the bottom, adjusting holder, put demo or user sample there, don't clamp it but let it rest (otherwise breaks when objective hits))

'''Brightfield''' (where does light come from, what do we expect to see)

'''Focusing''' (sample finding, and then moving around, warning: hitting lens to sample possible)

'''Epifluorescence''' (see visually light shining, see returning light, discuss different filters, user responsible for knowing their dyes and wavelengths, Searchlight spectra viewer)

'''Confocal scanning''' (gonna be a big part, mostly on computer now, switch in mood/user to focus on screen not on microscope - instructor sits on the far-right side of the desk to control the mouse, attendees sit right in front of the screen to see everything properly)

- Launch the software LAS X which controls the microscope, and choose the right startup settings (microscope = DMi8, settings = standard) and wait for it to appear completely. The software is going to ignore any wavelength/imaging modality (BF, EPI) choices you have made using the touch screen of the microscope and do confocal anyway.

- Overview of software areas:

# We start in "Acquire" (top) and will be spending most time there
# The big vertical line in the middle divides the view into settings (left) and image display (right)
# Making any adjustments to settings in the image display (right) is not going to change *any* raw data values or *any* microscope settings - just how the image looks on the screen
# Making adjustments on the left will change actual microscope settings, such as wavelengths used, scanning behavior etc.
# At the bottom, there are the buttons that make the microscope actually do something (Live, Capture Image, Start - etc.) - to prevent confusion when clicking "open laser shutter" in the software, or any other buttons.
# We make 1 small excursion to "Configuration" (left of "Acquire") and then "Laser Config" in order to switch on the lasers we know we will need.

- Laser setup

# choose the right wavelength
# give it a tiny bit of power (philosophy: start low, go higher if needed - instead of bleaching everything straight away)
# click round button under the laser to open the shutter
# Confirm yellow line appears in the schematic light path below -> laser ready.

- Detector setup

# Acknowledge we have a few detectors available, one is switched on.
# Prefer HyD (hybrid detectors), they are better. Typically they are in standard mode with 100 gain (digital gain) and you can use them like that.
# PMT detectors are less sensitive = good for strong signals (usually you have weak signals), to use one you have to give it some gain (couple hundred Volts, until image looks good to you)
# Move the detection window and resize it to capture the desired fluorescence emission wavelengths. Avoid including the laser wavelength at all cost!! This will visualise the laser, which we do not want to see - and it can damage the detector.

- First image

# Click "Capture Image"
# Probably, nothing will show up!
# Is there something there, but just weak? -> Set the Image Display (right) to "auto-scale" (button called "M" because it starts on Manual scale) -> have a look if something shows up.
# Is there nothing there anyway, maybe you need more laser -> Increase laser power to 20-30%, capture another image -> have a look if something shows up.
# Is there still nothing, maybe the focus is slightly wrong (we're in confocal mode after all...) -> use the Z-adjustment knob to focus up and down (either on the microscope, on the joystick, or on the USB panel) -> have a look if something shows up, now it really should.
# If there's still nothing, visually check if the sample is still alright, and then use Brightfield or Epifluorescence through the binoculars to find focus again. Then try confocal again, using the software

- Evaluating the first image

# It could be too weak (barely jumping out of the noise), so we need more laser until it's visible (improving the scan will help too, but this comes later) at the cost of possibly bleaching
# It could be too strong (huge areas look intense, and same intensity - mouse-over the pixel and see the maximum value 255 show up under the picture = too bright!) you want to avoid saturaiton

- Optimizing the scan (better picture, takes longer, make it as good as you need and not better)

- Switching to a higher magnification lens (use navigator to register 1 pic, switch to immersion lens, go back to original location).

- Z-stack (set start, end, use optimized)

- Time course (switch modes using dropdown menu, then set (1) interval (none, or amount of time) and (2) duration (unlimited, or amount of time, or amount of pictures)

- Tilescan (one small one, several of them, focus map)

'''Rounding off'''

- Saving data

- Saving settings

- Closing software

- Shutting down microscope (and cleaning oil objective, then put back at 10X)

- Leaving room and system tidy for next user

Leica SP8 Liachroic intro

2025-08-06T13:25:24Z

Jelle:

'''Intro-Intro'''

# Welcome, outline of the introduction
# The General Instrumentation department conceptually and physically
# The room and the general layout
# The microscope conceptually (functions)
# The microscope physically (components, roughly)

'''Switch-on'''

# Quick inspection (everything looks ok visually?)
# Switch-on procedure of components (buttons numbered order)
# Log-in in Windows (user will get an account) and software starting

Stage and objective movement controls (XY stage, Z objectives)

Sample placement (sample types, coverslip towards the bottom, adjusting holder, put demo or user sample there, don't clamp it but let it rest (otherwise breaks when objective hits))

Brightfield mode (where does light come from, what do we expect to see)

Focusing (sample finding, and then moving around, warning: hitting lens to sample possible)

Epifluorescence mode (see visually light shining, see returning light, discuss different filters, user responsible for knowing their dyes and wavelengths, Searchlight spectra viewer)

Confocal scanning mode (gonna be a big part, mostly on computer now, switch in mood/user focus on screen not on microscope)

Leica SP8 Liachroic intro

2025-08-06T13:05:08Z

Jelle: made a start

Leica SP8 Liachroic intro

2025-08-06T12:45:38Z

Jelle: created page

placeholder

Leica SP8 Liachroic

2025-08-06T12:45:19Z

Jelle:

Introduction protocol

Light microscopy

2025-08-06T12:45:08Z

Jelle: linked to sp8 liachroic

This page lists the options for performing light microscopy at the GI department.

=== List of equipment ===

==== Light microscopes ====

* Leica SP8X live-cell confocal
* [[Leica SP8 Liachroic|Leica SP8 liachroic confocal]]
* Leica "Thunder" widefield
* Leica "Timelapse" widefield
* PhaseView Alpha3 light sheet
* Leica DM2500 upright widefield

* [[Nikon TIRF widefield]]
* [[Nikon sora|Nikon SoRa spinning disc]]

==== Software packages and other tools ====

* Spinoza image analysis server
* Huygens
* Microfluidics pump for inverted microscopes

=== External links ===
[https://bookings.science.ru.nl/category/general-instrumentation-gi/gi-light-microscopy-image-analysis All microscopes (booking system)]

Leica SP8 Liachroic

2025-08-06T12:44:42Z

Jelle: created page

placeholder

Nikon sora functions

2025-07-28T12:52:21Z

Jelle:

== What can the Nikon SoRa do? ==
It can do live-cell, dual-color high-speed superresolution confocal imaging, in combination with FRAP

=== In more detail: ===

* Live-cell incubation (keeping temperature, humidity and CO2 at desired levels)

=== Observation by eye, using the binoculars: ===

* Brightfield (simple white light lamp, light coming from above the objective)
* Widefield epifluorsecence (specific excitation and emission colors, excitation light coming out of the objective)

=== Observation by camera, using the software: ===

* Brightfield
* Widefield epifluorescence
* Spinning disc confocal imaging (up to ca. 260 nm resolution)
* Superresolution spinning disc confocal imaging (up to ca. 120 nm resolution)
* FRAP (fluorescence recovery after photobleaching, 488 nm bleaching laser available)

Nikon sora

2025-07-28T12:52:06Z

Jelle:

[[Nikon sora functions|What can it do?]]

[[Nikon sora switch on|How to '''switch on''']]

[[Nikon sora superresolution switch|How to '''switch to superresolution''']]