Leica SP8 Confocal Microscope

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Dianwen Zhang (Email: [email protected])

Instructions for Leica SP8

Fluorescence Confocal Microscope Equipped with UV/Visible Lasers

Microscopy Suite Imaging Technology Group

Beckman Institute for Advanced Science and Technology University of Illinois at Urbana-Champaign

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405 North Mathews, Urbana IL 61801, USA http://itg.beckman.illinois.edu/ - I -

The Leica SP8 Confocal Microscope System: Overview

Figure 1 The system overview

While light source for bright field microscopy

Condenser Eyepiece

Transmitted Laser Detector (TLD PMT)

Front LCD control panel Scanner

System power console Mercury lamp for epifluorescence microscopy

Confocal imaging console

Focus knob Sample stage

Objective turret Stage controller –

Leica “SmartMove”

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COVID-19 Safety Standard Operating Procedures

This section was written to provide additional details to light microscope users to fulfill our online “COVID-19 Safety Standard Operating Procedures for the Microscopy Suite, Imaging Technology Group, Beckman Institute for Advanced Science and

Technology” (https://itg.beckman.illinois.edu/).

Pre-start checklist:

• Lab coat

Some of the supplies we normally provide may not have been replenished by the time you come in. It may be helpful to bring the following items with you:

• Small container of 70% alcohol

• Safety goggles

• A facemask

Disinfectant use:

Please consult with a staff member before using a disinfectant other than 70% alcohol (ethanol or isopropyl alcohol).

Never spray any disinfectant (including 70% alcohol) onto or near a microscope.

Disinfectant leaking into or dripping onto the optics, mechanisms, or electronic components of microscopes can cause costly damage.

Without confirmation or permission from a staff member, never wipe any parts of a microscope with disinfectant.

Be aware that many disinfectants (e.g. 70% ethanol) are flammable; please take

precautionary measures against the risk of fire from static discharge. This is particularly important when you are working on an optical table in a carpeted room. After disinfectant is applied, wait >30 seconds until 1) effective disinfection occurs and 2) flammable substances dry out, then 3) wave your hands to remove any vapors that may be floating around.

Lab coats:

If your lab coat comes into contact with surfaces that may be contaminated, stop work immediately and wash your lab coat or change to a new one.

Glove use:

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405 North Mathews, Urbana IL 61801, USA http://itg.beckman.illinois.edu/ - 3 - After touching any potentially contaminated surfaces with your gloves, spray disinfectant solution on them and wait long enough for effective disinfection to occur (usually more than 30 seconds) before continuing to work (refer to “Disinfectant use," above).

Lab safety goggles:

We encourage you to bring your own lab safety goggles.

Laser safety goggles:

If you are working in microscope rooms that require laser safety goggles, spray them with 70% ethanol (refer to “Disinfectant use, " above), and wipe them gently with Kimwipes until they are dry. The manufacturers of our laser safety goggles confirmed that 70% ethanol is safe for these products. Please do not use any other disinfectant.

Workspace preparation plans:

You should also receive a separate document that includes some examples of the use of the below listed plans on a few light microscopes in Microscopy Suite.

Plan A: For work surfaces such as benchtops or tabletops, it is acceptable to utilize

disinfectant solution or wipes. When spraying disinfectant solution on the surfaces, refer to

“Disinfectant use," above, and finish up by wiping any residues away with Kimwipes.

Plan B: For surfaces where disinfectant may not work effectively or may cause discomfort or damage (such as lab chairs, computer keyboards, computer mouses, joysticks, and most microscope parts, including focus knobs, eyepieces, touchscreen, etc.), please use the provided 18” wide plastic wrap to cover them up. See “Details of Plan B” below.

Note: Regarding microscopes, never place plastic wrap on lamp housings, any component that must be mobile, or any component with air vents.

After your use of the microscope, please remove all plastic film covers before you leave.

Plan C: For surfaces where disinfectant may cause costly damage and cannot be covered with plastic wrap (e.g. some components of a microscope, such as a motorized sample stage, objective lens turret, filter turret, etc.), we request that you touch them only with gloved hands. Refer to “Glove use," above.

Details of Plan B:

1. After entering the microscope room, clean your hands with hand sanitizer or soap and water if there is sink in the room.

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405 North Mathews, Urbana IL 61801, USA http://itg.beckman.illinois.edu/ - 4 - 2. Put on your lab coat and wear gloves. From this point on, wear your lab coat and gloves

until you have finished up and are leaving the lab. During any step listed below, if you accidentally touch a surface that is not disinfected, follow the “Lab coats" and “Glove use" policies.

Please follow the directions below, step by step, and repeat a step if you fail the first time. Some steps may be repeated as many times as necessary to complete them.

3. If the 18-inch-wide plastic wrap is mounted on a nearby wall rack, go to step 5; if not, find a work surface large enough (e.g. 18” x 36” for most computer keyboard + mouse) to permit Plan B to be carried out. Take a roll of the plastic wrap (caution: the outermost layer of the roll is potentially contaminated surface, so do not let it get contact with your lab coat) and lay it on the worktop. Spray the disinfectant solution onto the worktop and the plastic wrap roll. Refer to the “Disinfectant use" policy to assure safe disinfectant use and effective disinfection. You may need move the plastic wrap on the worktop a few times and turn it over to make sure that all surfaces (including your gloves) are exposed to disinfectant and thus become disinfected.

4. Unwrap the roll*. Patience and practice are needed to keep the unwrapped film from folding and sticking to itself. Once a piece of film has been unwrapped to a suitable length, cut it with scissors and pick it up carefully. Go to step 6.

5. Pull down the outermost layer of the roll from the wall rack, cut with scissors, and discard the part that had been exposed, as it is potentially contaminated. Spray disinfectant solution on your gloves and the scissors, and wait for disinfection to complete. At this point, you can now pull the plastic wrap off to the desired length and cut it with scissors. You may need to use your other hand to support the film and keep it from folding in on itself.

6. Carefully hold the film until you can lay it completely onto the surface that you want to cover*. Check to make sure that the film can cover the entire surface area (including, e.g., the edges of the computer table and chair).

7. Repeat steps 3 through 6 until you have covered all surfaces you may contact during your experiment.

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The demo video is available at https://youtu.be/froBmoIP7eE for steps 4 and

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An Example of the Working Space Preparation in the Microscope Room for COVID-19 Safety

In the below photo (Figure 2), I demonstrate an example of my practice for the

abovementioned COVID-19 safety SOP. It is, of course, in no way exhaustive. You should adapt it to your needs in your experiment and make sure every surface which you can come into contact with is prepared to work with it safely.

Figure 2 An example working space preparation for COVID-19 Safety Plan B

Plan C

Do NOT cover the lamp house

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Remote Training or Technical Assistance

Please note that a webcam with built-in microphone is available in the microscope room for remote training or technical assistance. You may unplug it from the PC or place it facing down on the table to ameliorate any privacy concerns.

Use “Skype for Business” to initiate contact

You can find the software “Skype for Business” on your desktop. Click to run it. After its front panel is open, please search (see the example screenshot in Figure 3, pointed by the red arrow) for my phone number 217-333-4387 or my email [email protected] to find me.

Figure 3

After you see the listing picture for me, move the mouse onto the picture (see the example in Figure 4, pointed by the red arrow) to see your options (such as message, voice call or video call). You can usually start from text messaging me (pointed by the blue arrow in Figure 4).

Figure 4

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Remote visual inspection or desktop control

In case of online training or remote technical assistance, remote visual inspection and desktop control are needed. Please find the webcam, and plug it to the computer when you are ready for using it.

First, you should follow the instruction in the previous section (titled Use “Skype for Business” to initiate contact) to initiate contact with me. If I am available, you can start the

software Zoom by clicking on the icon on the computer desktop. When the Zoom window is shown up, choose “Join a Meeting” (see in the example screenshot in Figure 5, pointed by the red arrow).

Figure 5

You will see the below window, and then please wait me for a few minutes. I need set up to host a zoom meeting, and give you the meeting ID and the password as soon as I have them.

Figure 6

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Start Up the Microscope System

Figure 7

Following the numbers in Figure 7, push the green buttons to turn on the system powers in the following order (from left to right): the microscope ① , the scanner ②, the lasers power ③ (turn on the laser key ④) and the mercury lamp ⑤ (Check the black button ⑥ which should always be in

“Remote” position).

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Start Up the Software LAS X

1. Make sure that there is no sample loaded onto the stage, and then start the control

software “LAS X” .

Keep all default settings in the configuration startup window (Figure 8), click “OK”.

Figure 8

It will take 1-2 minutes for the software to find and initialize all components in the microscope system, before the below main window (Figure 9) of the software is up.

Figure 9

2. By default, LAS X starts with the “TCS SP8” module ①, and opens the “Acquire” tab window ② (see Figure 10). Please do NOT change these settings unless you are trained to do so. Note: Changing the settings in the “Configure” tab window ③ can cause hardware failure or damage.

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405 North Mathews, Urbana IL 61801, USA http://itg.beckman.illinois.edu/ - 10 - Figure 10

3. Some commonly used settings for confocal imaging are highlighted in Figure 11.

Figure 11 Typical scanning speed is 600 lines per second ①.

When turning on “Bidirectional X” ②, “Phase X” ③ will show up. Based on some other settings, LAS X automatically modifies the value of “Phase X”. Manually changing this parameter may cause uneven lines in confocal image.

4. Change the laser intensity control panel user interface (UI) to classical UI.

Figure 12

Click the “Switch to Classical UI” button (pointed by yellow arrow in Figure 12) to use the classical UI (Figure 13) showing clearly all laser lines that are available in the system.

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405 North Mathews, Urbana IL 61801, USA http://itg.beckman.illinois.edu/ - 11 - Figure 13

Turn on the “ON/OFF” buttons (pointed by cyan arrows in Figure 13), respectively, to enable the UV or Visible laser lines control panel or both.

5. Enable the laser lines that you need

Click anyone of the two “+” buttons (pointed by yellow arrows in Figure 13) to open the below window (Figure 14).

Figure 14

Make sure that both “UV” and “Visible” buttons (pointed by yellow arrows in Figure 14) are activated (in red) to see all laser lines.

When turning on the “Argon” laser to enable 458 nm, 488 nm and 514 nm laser lines, set the output power to 20% (pointed by the cyan arrow in Figure 14) of its maximum power.

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405 North Mathews, Urbana IL 61801, USA http://itg.beckman.illinois.edu/ - 12 - 6. Enable the detectors/channels for your fluorophores

There are three detectors, PMT 1, PMT 2 and HyD 3 (higher sensitivity), in the system.

Therefore, the system can support up-to-three-channels simultaneous imaging.

Click the ON/OFF buttons (pointed by cyan arrow in Figure 15) to enable the detectors that you need.

Figure 15

The color of display output of each channel can be set from a pull-down menu by

clicking the round colored button (pointed by yellow arrow in Figure 15) on the left side of the name of each detector.

There is the fourth detector, TLD (Transmitted Light Detector), for bright field imaging with the laser light transmitted through sample. To use this detector, click the triangle button (pointed by red arrow in Figure 15) to show the control panel.

7. Set up the detection spectral range

Figure 16

The detection spectral range can be modified in either direction by simply clicking on and dragging the edges (pointed by yellow arrows in Figure 16) to match the emission spectrum of the fluorophore assigned to the channel.

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405 North Mathews, Urbana IL 61801, USA http://itg.beckman.illinois.edu/ - 13 - 8. Choose objective lens

Figure 17

Objective lens can be changed either from the “Objective” pull-down menu in the software or manually by turning the objective turret on the microscope stand.

To check the specification of each objective lens, click the button indicated by the cyan arrow in Figure 17.

After setting up the software, you can load your sample and find it under the microscope in white light bright field microscopy or epifluorescence microscopy.

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Find Sample through the Eyepiece

After a sample is loaded onto the microscope stage, use the second tab (pointed by green arrow in Figure 18) from the front LCD control panel of the microscope to select proper microscopy function for sample focusing through eyepiece.

Do NOT change any settings in other tab windows.

Figure 18 The front LCD control panel of the microscope

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Bright field (BF) microscopy

Following the numbers in Figure 19, finish the below steps to start BF microscopy for sample focusing:

1. Switch to BF microscopy ① and open the “TL-Shutter” ② from the front LCD control panel of the microscope.

2. Adjust the illumination intensity by turning the “TL/Fluor” dial ③ located near the focus knob on the left side of the microscope stand.

Figure 19 1

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Epifluorescence microscopy

Following the numbers in Figure 20, finish the below steps to start epifluorescence microscopy for sample focusing:

1. Click the “FLUO” button ① on the front LCD control panel of the microscope to switch to epifluorescence microscopy.

2. Choose right fluorescent filter set ② (the three buttons, DAPI_LP, FITC_LP and

RHOD_LP, are for ultraviolet, blue, and green excitation, respectively. They are simply longpass bandpass filters).

3. Click the “IL-Shutter” ③ button to open the shutter.

4. Adjust the illumination intensity by turning either the “Intensity” dial ④ on the mercury lamp or the “TL/Fluor” dial (see it in Figure 19) located near the focus knob on the left side of the microscope stand.

5. Perform focusing. Move the sample into the focus of the objective lens.

6. Click the “IL-Shutter” ③ button again to close the shutter to avoid fluorescence

photobleaching. Note: if this button cannot close the shutter, check and make sure that the black button at the left bottom corner on the front panel of the mercury lamp is in

“Remote” position.

Figure 20

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Confocal Imaging

The microscope condenser has a laser safety interlock at its bottom to assure no laser can go through the system when you push back the condenser to load sample. After you load sample, make sure to pull the condenser forward to its normal upright position to disengage the interlock for confocal imaging. If you forget, LAS X will show you an error message saying that the interlock is open.

Live imaging

1. Find sample through microscope eyepiece in either bright field microscopy or epifluorescence microscopy (refer to the previous sections).

2. Click the button at the left bottom corner of LAS X to start live imaging. On the right side of the software, image window for each channel/detector that you activate to use (refer to the previous section “Start Up the Software LAS X”) will show up.

3. Click on an image window to select the detector. The image window will be highlighted with a white line square box. Follow the below steps to get the first image:

1) Start with no laser on the sample (all laser intensities are 0%). Set the gain of the detector to highest possible (the maximum values are 1250 v for the PMTs, and 500%

for the HyD, respectively) either from the software or by turning the leftmost knob,

“Smart Gain” on the “confocal imaging console” (Figure 21) right in front of the computer monitor.

Figure 21 Confocal imaging console

2) Gradually increase the laser intensity by dragging the laser power bar until the image appears on the image panel. However, if you fail to see the image, even when the laser intensity is set very high (e.g., > 80%), stop live imaging and go back the microscope stand to check objective focus through eyepiece (refer to the previous sections for bright field microscopy or epifluorescence microscopy).

3) Turn the rightmost knob “Z Position” on the “confocal imaging console” clockwise and/or counter-clockwise in turn to move the sample up and down within 500 µm

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405 North Mathews, Urbana IL 61801, USA http://itg.beckman.illinois.edu/ - 18 - relative to the objective lens until a good image is found.

4) Balance the detector gain and the laser intensity to get sharpest image. High

detector gain causes high background noise, so lower detector gain is desired, but it will require increasing laser intensity, while high laser intensity can cause sample fast photobleaching. Practice to find the balance in between these two control parameters for the best image.

4. Repeat step 3 for all image channels. Figure 22 shows an example from a three- fluorophores-stained sample. Click on the “Overlap” button ① on the right side of the images window to see the fourth three-channel-overlapped image at the right bottom corner.

Figure 22

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2D imaging

The “Capture Image” button at the bottom of LAS X is only for two-dimensional image acquisition from the current objective focus.

3D imaging (Z–Stack)

Figure 23

1. Click the button at the left bottom corner of LAS X to start live imaging.

2. Turn the rightmost knob “Z Position” on the “confocal imaging console” (Figure 21) to one direction all the way to one surface of the sample volume that you intend to image, and then click the “Begin” button ① in the “Z–Stack” control panel (Figure 23).

3. Now turn the “Z Position” knob back all the way to the other surface of the sample volume, and then click the “End” button ② in the “Z–Stack” control panel.

4. If you want to control “Number of Steps” or “Z-Step Size”, you can manually type in ③, or usually you can stay with the default setting “System Optimized”.

5. Click the button to start Z–Stack acquisition.

6. When the settings in the “Z–Stack” control panel are not needed after the acquisition is complete or aborted, delete them by clicking on ④. Or 3D imaging will be run whenever the button is hit (for example, for tile scan imaging in the next section).

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Tile Scan for large image

1. Activate “Tile Scan” (pointed by cyan arrow in Figure 24) from the “Acquisition Mode”

Figure 24 menu, and then the “Stage” control panel (Figure 25) will emerge at the left side of LAS X.

Set the “Zoom Factor” (pointed by yellow arrow in Figure 24) to 1.2 for scan imaging. When “Zoom Factor” is too small, the laser beam needs to be steered to a big angle away from the optical axis to image a large area, and thus it may partly hit on the back aperture of the objective lens, causing weaker intensity at the image edges. This can leave visible seam between any two neighboring images after stitching.

2. Set “Zoom” ① (Figure 25) for the stage map in the

“Stage” control panel to make the map able to cover the entire area that you intend to image with the tile scan.

3. Start live imaging by clicking the button at the left bottom corner of LAS X.

4. While observing the live image window, move your sample with the stage controller (or called

SmartMove by Leica. Note: You may need to change it to “Precise” mode first) to find a corner of the area to be tiled, and then ② click to register this position.

5. Repeat step 4 to register as many corner points as you need until the final grid in the stage map can cover the entire area. Double-click the mouse on the grid, you can check the live image at the point.

6. Turn on the “Merge Images” button ③.

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Figure 25

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405 North Mathews, Urbana IL 61801, USA http://itg.beckman.illinois.edu/ - 21 - 7. Choose “Overlap blending” ④. Typically, “Smooth” is often used.

8. Set Speed / Accuracy ⑤.

9. Click the button to start tile scan. Note: if you have undeleted “Z-Stack”

settings, tile scan will run in three dimensions.

10. When acquisition is complete, the images will be automatically stitched together to form a composite image.

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Sequential scan

Figure 26 shows an example of how “sequential scan” is usually set up to use.

1. Click the rightmost button ① in the “Acquisition Mode” menu to enable/disable

“Sequential Scan”.

2. Sequence change can be set ② in “Between Lines”, “Between Frame” and “Between Stacks”. Note: imaging speed can be significantly reduced when changing sequence

“Between Lines” if every sequence uses different laser line and/or has different detection spectral range. “Between Frame” should be usually used.

3. Click the “–” or “+” button ③ to remove or add a sequence. In the example, two more sequences, “Seq. 2” and “Seq. 3”, are added.

Figure 26

4. Double-click on a sequence button ④ to activate and set up the sequence.

5. Choose a detector and set its detection spectral range ⑤.

6. Follow the instruction in the previous section “Live imaging” to set the laser intensity and the detector gain for the selected sequence.

7. Repeat steps 4-6 for all sequences.

8. Click the button to start sequential scan. Note: if you have undeleted “Z–Stack”

settings, sequential scan will run in three dimensions.

Note: The settings for sequential scan won’t be saved to data file, so if you want to reuse the 1

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405 North Mathews, Urbana IL 61801, USA http://itg.beckman.illinois.edu/ - 23 - settings, please click the “Save” button ⑦ to save it and load it next time when you need it.

Save and Post-process Data

Open the “Open projects” tab (Figure 27) on the left side of LAS X, you can find a list of your image data. You can save them to a single project file *.lif together with the software settings for each acquisition (except for “Sequential scan”).

The free software Fiji (https://fiji.sc/) can open *.lif file directly and has a great deal of functions for post-processing fluorescence confocal image data.

Figure 27

Import Previous Settings from Project Data File

To reuse the software settings from a previous image data, open the project file, and click to select a favored image or images set, and then click the “Apply” button (pointed by yellow arrow in Figure 27). The acquisition settings (including laser intensities, detector gains, detection spectral ranges, etc.) for the image or images set will be applied to the system control panel in LAS X.

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Shut Down the Microscope System

Figure 28 1. Turn off the mercury lamp ①.

2. Close LAS X.

3. Wait until LAS X completely exits to turn off the system powers in the following order: “PC/Microscope” ②, “Scanner Power” ③, the laser key ④, and then wait 5 minutes to turn off the “Laser Power” ⑤.

Do NOT turn off the laser power immediately after turning off the laser key. If you accidently make this mistake, immediately turn the laser power back on to allow the laser cooling down.

4. Log off from the computer (Do NOT shut it down).

5. Use Kimwipes to wipe off immersion oil from objective lenses.

6. Clean up the working place.

1) Remove all plastic films that you used to cover the workspace.

2) Used glass slides or coverslips should be thrown into sharps waste bin.

3) Take away all your stuff.

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