IGB Core Training
This book will contain Training materials for each instrument

600MHz NMR
NMR Training

Agilent Hardware


Basic Operation
600 MHz NMR 

 “In the absence of a magnetic field the period of all these oscillations is the same. But as soon as the electron is exposed to the effect of a magnetic field, its motion changes.” – Zeeman, P. 

 FIRST PROCEDURES 

 Content: 1. Sample preparation 2. Logging in 3. NMR instrument 4. Some important line commands 5. Data acquisition      5.1. Locking      5.2. Shim 6. Annexes 7. References 8. Troubleshooting 

 1. Sample preparation 

 “There are no good results using bad samples.” 

 CAUTION! Clean the sample of precipitates, cells debris, bubbles, foam, any solid and heterogeneous phases at all in your sample, because different phases will cause distortion on the field and will impact the result. To deal with this issue, use ultracentrifuge in a highest speed as possible to deposit any solid, and/or filter the sample using at least 0,22 µm pore size filter (attention to the filter composition to avoid dissolution of the material by the solvent) right before fill the NMR tube. This step is crucial to obtain a well-adjusted shimming applied to your experiment. 

 ALAWAYS! Clean and dry the sample tube. A tiny scratch, humidity, oily, or dust can destroy the high quality properties of the tube making it useless for high resolution experiments, and also leaving dirty to the inside of the probe, reducing the sensitivity and harming the magnet. 

 Make your sample using a deuterated solvent (often deuterated water is used for polar samples at 10-20% v/v) until the total volume reaches a volume of ~ 500 to 600 µL (or 0.5 – 0.6mL) in a 5-mm tube, 3.1 mL in a 10-mm tube, and 150 µL (or 0,15 mL) for a Shigemi® tube are adequate for removing the end effects. In general, the NMR samples are compatible with: Acetone-d6, chloroform-d, methylene chloride-d2, and DMSO-d6. Make sure that the solvent will not react or degrade the sample and has a high purity for NMR, free of any impurity. Remember, the peaks of the impurity will appear in the same spectrum of your sample, which can cause a misinterpretation of the result and loss of the sensibility (gain). 

 If the impurity is part of the solvent, is possible to dry the contaminated solvent through N2 stream and dissolving the sample in an appropriate solvent again (repeat the process if you observe any improvement on the result, usually monitored by 1D NMR screening is enough. 

 The liquid column length must be at least three times the length of the observe coil window to minimize end effects (fig. 1). A typical sample length is 5 cm regardless of the diameter of the NMR tube used, positioned properly. Make every sample up to the same height in order to obtain similar shim values using samples of that height. 

 ATTENTION! Some probes may require particular volume and height adjustments. Reduction of sample volume to attain higher concentration usually fails, at this case some special plugs for low volume samples are available and will help with line shape. As well the temperature. Leave the sample become in thermal equilibrium with the probe surrounds, the suggested procedure is to wait for at least 5 minutes or the VT panel/monitor shows a stable behavior with the sample tube inside of the probe before to start any intervention on the equipment, is known that a difference of 1oC can affect the chemical shift in 0.01 ppm. 

   

 

 Figure 1. Sample depth gauge: 

 As a final point, there are few questions to do just before you perform the NMR experiment, as follows: 

 1. Do I used the depth gauge properly? 2. Do I filled the NMR tube with sufficient volume? The tube is under filled or overfilled, which might cause shim problems? 3. Is the sample homogenous? 4. Are there some particles suspended or precipitated? 6. Are there some paramagnetic substance or high ionic concentration? 7. Is there any possible intrinsic chemical dynamic or exchange in some of the components of my sample? 

 If your answer was yes for any of these questions, the standard parameters will be modified caused by this specific environment, consider go back to fix any inconvenient, if not possible, consider this specificity during the whole procedure. 

   

 2. Logging in “Last night I slept like a log. I woke up in the fire place.” – Cooper T. 

   

 2.1. Lock 

   

 2.2. Shim 

   

 3. NMR instrument “It is the photographer, not the camera, which is the instrument.” – Ardold, E. 

   

 

 Figure XX:  

   

   

   

   

   

      3.1. Locking 

   

      3.2. Shim 

   

   

 4. Some important line commands “No command, no work.” 

   

 5. Data acquisition “Experts often possess more data than judgment.” - Powel, C. 

 Always remember because these experiments involve a fourier transform that to double your signal-tonoise ratio requires four times as many scans. So if you can’t see anything after 1h you are not going to have much to look at after 4h, likewise if you can not see anything after 2h it is not going to improve a lot after 8h. So on long experiments check your data after an hour or so and if can not see any correlations you might as well stop the experiment. 

   

   

      5.1. Locking 

   

      5.2. Shim 

 When the magnet shims are not optimized, the lines in your NMR spectra can take on distinct shapes. The shape of the peak will give you a hint as to which shims need adjustment. Note that lineshapes can also be much more complicated than those below and can even show splittings. 

   

 6. Annexes “The reward for work well done is the opportunity to do more” – Salk, J. 

   

 7. References “Everything I do references something that influenced me” – Abloh, V. 

 [1] NMR Spectroscopy, User Guide Varian, Inc. Inova and MercuryPlus NMR Systems With VnmrJ 2.2MI Software Pub. No. 01-999378-00, Rev. A 0708 

   

   

   

 8. Troubleshooting “It's so much easier to suggest solutions when you don't know too much about the problem.” – Forbes, M. 

 

   

  

NMR Theory


Axio Scan Z1
Axio Scan Z1 Operating Manual 

 1. Make sure the slide scanner switch on the back and on the front 

 2. Login to the computer with your NETID and IGB password 

 3. Start ZEN blue then clicks on ZEN slidescan 

 4. Load your slides into the trays, place your slides face up with the label in between holder then press the silver button on the instrument or open using Zen.  

 

 Note: occupied slots are highlighted on slide scanner in blue, green, orange, red or white. 

 Blue = tray loaded and no slide preview taken. Slide status is “new”. 

 Green = tray loaded and slide status is either “preview” or “finished”. 

 Orange = an error occurred during scanning or tray loading. 

 Red = probably a tray is stacked inside the slide scanner. 

 White = tray is inserted but empty 

 5. Storage location: direct the folder to save your images in: D:\User Files Deleted Anytime  

 Note: After scanning, make sure you transfer your data to the core server under your PI’s NETID: \\core-server.igb.illinois.edu   

 6. Mark All : All slides are displayed in the Magazine tab that appears automatically when slides are loaded into AxioScan.Z1. Unmark All slides to avoid compromising other experiments. 

 7. Select profile from default scan profile then click the gear icon to open advanced wizard and modify your settings 

 Setting Up Brightfield Scan Profile   

 1. Global Data 

 Profile Type is where you define if you are going to scan in brightfield or fluorescence 

 Tissue Detection Mode should be in principle always Automatic , because compared to Interactive and Manually (the defined regions will always be the same for all slides assigned with this profile) , the preview is done immediately and not after editing the tissue detection. 

 TD Recognition Type for brightfield profiles should only be set to Tissue (via thresholds). This means that detection will be based on the RGB colors of your slide’s preview, compared to Marker where detection is based on the recognition of a pen contouring your section, this option is only recommended for fluorescently labeled sections, which are not visible with Preview Cam. 

 

 Label Scan Settings : This tab allows you to adjust the size of the window displayed in red that defines the slide label image size and position, the image always save in separated file called pt1 . 

 Preview Scan Settings : the red rectangle defines the space on the slide that will be used to detect the edges of tissue detection. Note: Sometimes it is useful to reduce this area to improve the automatic tissue recognition on the slide borders, you might want to check if the predefined size corresponds to your needs. The image always saves in separated file called pt2 .                

 Tissue Detection Settings: check Automatic and use Test to check the quality of the recognition. The display curve ( C ) can be modified to change the contrast of the image (set as best fit) and help your eyes to see the tissue clearly. The yellow ROI shows where Zen will obtain data for the shading correction. Notes: Green ROIs are defined by the following parameters (see Application guide AxioScan.Z1 v1.1 ):

 4.A. Specimen histogram ( A ) can be modified to select specific intensity values (detail in fig below). 

4.B. Region dilation size ( B ) defines a value for spacing the border around the detected tissue. 

 

 

 5. Focus Map Settings: Axio ascan Z1 uses a two-step focusing method ( Coarse and Fine ) to give the best results. 

 5.1. Coarse focus looks for tissue’s macroflatness, which basically means that tries to locate the section on the slide. 

 lower magnification objective (5x) is recommended to generate coarse focus                

 Navigate to the specimen with the tool you have in the lower part window. You can click inside the green ROI where do you want to focus or use virtual joystick within stage tab.                                              

 The next step is to focus on the specimens and adjust both exposure time and light sources intensities or click measure to adjust automatically.                                                                                                                                                                      

 Activate Run Auto Focus to see a live image from camera and adjust focus automatically. If the autofocus fails, focus can be adjusted manually using expander Focus. 400-600 µm range of focus is recommended (the higher the range the longer it takes to find focus). For 5X focus is typically around ~4000µm (look up current z position, in the image below the focus at 4120 µm then manually set first= position-200 µm and set last=position +200 µm) to make sure that the focus on the center of the range.                                                                                                                                                                                

 Focus point distribution strategy: using Number of points (parameter=4-12) is the default setting to get the best value of wide variety specimens. To change the focus point distribution strategy set, double click on the entry. A window comes up, you can select other strategy and click on modify the changes will be applied. Note: Detail explanation of focus strategy for different type of tissue can be found on Application guide AxioScan.Z1 v1.1 pg 26-28 . Some trial and error are always needed to fine the right compromise between speed and focus quality, in general, the flatter the sample is the better.    

 

 5.2 Fine focus looks for tissue’s microflatness which identify small focus within the section. 

 Use the same objective for fine focus as when you scan the slide                                                                                                

 

 Click Run Auto Focus to perform autofocus, or using Focus expander to adjust focus manually, in this step adjust the exposure time and/or intensity of light source to get a signal around 5% of dynamic range so you can reduce time for focusing and reduce photobleaching of the sample.                                                                                                  

 

 Range coverage defines whether the complete z-stack is performed (Full) or if it is only captured until local maximum is detected first (smart). Full is more robust while smart is fastest. We recommend range of focus 50 to 100 µm (max). 

 Focus point distribution strategy set : Onion skin is the standard setting for the fine focus map, providing an even distribution and ensuring the border has enough focus point, Onion skin 0.1 means that 10% of the number viewing files will be used as focus points to calculate the focus surface. Note: Onion skin strategy is particularly useful for medium and irregular tissue section under fine focus by creating concentric ring within tissue detection. Center of gravity is recommended for Tissue micro arrays. Detail of Focus point distribution strategy can be found at Application guide AxioScan.Z1 v1.1 .                                                                                                                                                

 6. Scan Settings : Adjust the flash intensity or flash duration to get desire image for final set up.                                                                    

 

 Online processing: the standard setting is online setting as, the pyramid active should always be activated, as this will speed up the viewing the image afterwards. The final setting allows to adjust the compression which be applied as part of online processing. JpegXR active is activated via checkbox, you can save lossless compression which no information is lost If lossy compression is activated the quality should set to 85%.                                                                                  

 Click finish                                                                                                                                                                          

 Save the profile using Save adapted scan profile and make sure to change the scan profile to the new scan profile             

 

 Start Preview Scan : this setting generates the preview that will be used as defined by the assigned profile, it consists of label area and specimen area.                                                                                                                                                                                                                                                  

 

 Start Scan : to image the slide. 

 To open the image, double click on it and it will separate open in a separate tab in the center of the screen.            

 Fluorescence Scan Profile 

 1. Global data: the profile type should be fluorescence, tissue detection mode is automatic, TD recognition type for fluorescence is marker. Channel is recommended mode for AF contrast type coarse or AF contrast type fine                                                      

 Note AF Contrast Type Coarse and Fine are channel and RAC, only for fluorescence profile.  

 Channel: use fluorescence signal to perform focus with one of your florescence stains (preferably a counterstain), you must ensure that the stain evenly distributed over the sample, so the system detects enough signals at the focus point to preform reliable autofocus 

 RAC: system use ring aperture contrast to perform, this recommended if no fluorescent counterstain is available and/or if fluorophore is very sensitive to bleaching. 

 2. Label Scan Setting                                                                                                                                                                                                                                                                      

 

 3. Preview Scan                                                  

 4. Tissue detection settings: check automatic and test, if green ROI does not cover your tissue, you can adjust the histogram under marker (A), region of shrink size (B), adjust the histogram on the display to best fit to see the tissue clearly (C).    

 5. Focus Map Settings: Axio scan Z1 uses a two-step focusing method ( Coarse and Fine ) to give the best result. 

 5.1. Coarse Focus 

 Lower magnification objective (5x) is recommended to generate coarse focus 

 Choose 1 channel/fluorophore that bright and uniform on your tissue, DAPI is typically recommended as the channel to use for both coarse and fine focus. If your sample has absolutely no signal in DAPI you may use a different channel for focus reference. 

 Navigate to the specimen with the tool you have in the lower part window. You can click inside the green ROI where do you want to focus or use virtual joystick within stage tab. 

 The next step is to focus on the specimens and adjust both exposure time and light sources intensities, mainly adjust the exposure time to get the intensity histogram around 1000, this value is enough for focusing, which allow speed up the focus map and lower photobleaching of the sample. 

 Adjust focus automatically using Find focus/AF or you can adjust manually using expander Focus if the autofocus fails. Activate Live to see a live image from camera                                                                                                                  

 Focus point distribution strategy: using Number of points (parameter=4-12) is the default setting to get the best value of wide variety specimens. To change the focus point distribution strategy set, double click on the entry. A window comes up, you can select other strategy and click on modify the changes will be applied. Note: Detail explanation of focus strategy for different type of tissue can be found on Application guide AxioScan.Z1 v1.1 pg 26-28 . Some trial and error are always needed to fine the right compromise between speed and focus quality, the flatter the sample is the better. 

 5.2. Fine Focus 

 Fine focus looks for tissue’s microflatness (small focus variation within the tissue) 

 Use the same objective for fine focus as when you scan the slide for example 20X objective (minimal) 

 Click Find focus (AF) to perform autofocus, or using Focus expander to adjust focus manually, in this step adjust the exposure time and/or intensity of light source to get a signal around 5% of dynamic range so you can reduce time for focusing and reduce photobleaching of the sample. 

 Software autofocus: the setting under quality defines the type autofocus algorithm in use. The default setting is Quality is best and Range coverage is Full. Range coverage defines whether the complete z-stack is performed (Full) or if it is only captured until local maximum is detected first (smart). Full is more robust while smart is fastest. 

 We recommend a range of focus 50 to 100 µm (max). 

 Focus point strategy : Onion skin is the standard setting for the fine focus map, providing an even distribution and ensuring the border has enough focus point, Onion skin 0.1 means that 10% of the number viewing files will be used as focus points to calculate the focus surface. Onion skin strategy is particularly useful for medium and irregular tissue section under fine focus by creating concentric ring within tissue detection. Center of gravity is recommended for Tissue micro arrays. N umber of points =4 for small tissue and N umber of points = 6-8 points for medium size number of points 6-8 points. Note Focus point distribution strategy (see Application guide AxioScan.Z1 v1.1 )                                                

 6. Scan settings: 

 Add a new channel by clicking the plus sign, and choose dyes from the list, you should check that the light path if the correct filter is chosen for each fluorophore. 

 Always check the tissue in a spot where you expect to have higher expression so that you can adjust exposure time to avoid saturation of the signal. 

 Click find focus/AF: find the focus automatically, all the channels are displayed in the image and intensity curves are shown in the histogram 

 Adjust exposure time and lamp intensity of each channel to try and use about 50% of the dynamic range.          

 Online processing: the standard setting is online setting as, the pyramid active should always be activated, as this will speed up the viewing the image afterwards. The final setting allows to adjust the compression which be applied as part of online processing. JpegXR active is activated via checkbox, you can save lossless compression which no information is lost If lossy compression is activated the compression.                                                                                              

 Click finish                                                                                                                                                            

 Click the Gear Icon and save the profile using Save adapted scan profile and make sure to change the scan profile to the new scan profile                                                                                                                    

 Start Preview Scan: this setting generates the preview will be used as defined by the assigned profile, it consists of label area and specimen area.                                                                                                                  

 Start Scan: begins to image the slide. 

 To open the finished image, double click on it and it will separate open in a separate tab in the center of the screen. 

 Tutorial videos on how to set up brightfield and fluorescence for Axio Scan Z1 by Zeiss can be found in this link https://www.zeiss.com/microscopy/us/local/zen-knowledge-base/axio-scan.html 

   

   

   

   

   

   

   

   

  

Axio Zoom V16 Operating Manual
Power ON Routine 

 1. Turn on X-Cite epi-fluorescence light source for fluorescence imaging 

 2. Turn on EMS3 box 

 

 3. On the SYCOP 3, the system will prompt you to calibrate the stage: click yes to perform stage calibration. Make sure there is nothing blocking the stage. 

 4. Please check that you have no sample on the stage prior to running calibration. Click ‘OK’, The stage will move to the extreme ends of its travel range during the calibration, it is important that no sample be mounted to avoid any potential damage to the sample and/or the system.                                                                                                                  

 5. Login to the computer Using NetID and IGB password 

 6. Double click on Zen blue software then click Zen Pro and wait until initiation process is completed. 

 7. Click calibrate now then wait for the SYCOP to complete the focus calibration before mounting sample.                            

 On the Zen software:                                                                                                                        

 A. Image Acquisition Tools Represented by a group of blue bars each containing a series of tools for sample observation (Locate), image acquisition (acquisition), image processing and system maintenance . Locate tab also contains primary image capture functions. For imaging we will use locate and acquisition tab. 

 B. Image Display This area is where each newly captured image will appear. Settings here allow the user to control how the image is viewed on screen. 

 C. Catalog of Open Images This list displays each image that is currently open within the ZEN software. This area contains tools for saving data sets. 

 Mounting Sample 

 1. Place sample (in the center of field of view) underneath microscope objective. 

 2. Push eyepiece plunger IN to send light to eyepieces.                                                                                                                        

 Fluorescence Imaging using AxioZoom V16   

 3. From Locate tab select filter for fluorescence (DAPI, FITC, AF 568) and reflected light ON                                  

 4. Using stage and focus controls, position (X, Y) and focus Z via the eyepieces                                              

 5. When a suitable location has been found, pull the eyepiece plunger OUT to camera 

 6. pull the mirror, for fluorescence we used AxioCam HR, 100% transmittance, 0% reflected, 

 7. Switch from locate tab to acquisition tab                                                                                                        

 8. From Channel dialog, add track, dye, ref color, and camera (AxioCam HR) 

 9. Highlight the track and click set exposure. Note: Set Exposure is an automatic exposure measurement taken directly from the area of the sample exposed. If you have reasonably bright fluorescence, then this function works well but it may need to be fine-tuned. If your exposures are dimmer than required, you can either manually increase the exposure ‘Time’ slider or increase the ‘Shift %’ and run Set Exposure again. If your exposures are brighter than required, you can either manually decrease the exposure ‘Time’ slider or decrease the ‘Shift %’ and run Set Exposure again. Or click Live to adjust exposure time. You should use the range indicator to check if you have saturation in your image then adjust exposure time and shift.                            

 10. Repeat steps 9 for each Track (or color) you have. 

 11. When the exposure time has been measured for all Tracks, click ‘Snap’ to collect an image with all colors. 

 12. Select save the image 

 Bright field imaging using AxioZoom V16 

 1. Click Locate to see sample using the eyepiece, choose empty filter (4), make sure the light into eyepieces by pushing eyepiece plunger IN to send light to eyepieces.                                                                                                                      

 2. The brightfield light source is from the bottom or top (turn on the switch for the top LED light source)                  

 3. When a suitable location has been found, pull the eyepiece plunger OUT to camera                              

 4. push the mirror, for brightfield we use AxioCam color (100% transmittance, 0% reflected)                

 5. Switch from locate tab to acquisition tab 

 6. From Channel dialog, add track, choose TL brightfield, and color camera (AxioCam 512) 

 7. Highlight the track and click live and adjust exposure time. You should use the range indicator to check if you have saturation in your image then adjust exposure time and shift. On the display histogram you can check how much of the dynamic range you are using.                                                                                                                                                      

 8. From the Acquisition Mode dialog, click ‘Auto’ from the White Balance settings. Note: The ‘Auto’ feature works reasonably well on most samples. You may need to use the ‘Color Temperature’ slider to optimize the image. 

 9. Click snap for a final image 

 10. Save the image 

 Using AxioZoom for Tiling 

 1. Check ‘Tiles’ from the experiment options. This will activate the Tiles dialog in the Multidimensional Acquisition settings 

 2. From the Tiles dialog, click ‘Advanced Setup’ to begin defining your tile regions. You can choose how many tiles and X and Y and click the plus sign to add the tile region. Alternatively, click Advanced Setup, this will open a preview window that allows you to interactively define your tile regions from the Live image display. The idea here is that we will define the tile boundaries from which ZEN will calculate the number of tiles required to cover the area selected 

 3. Using the XY joystick, move your stage position to the left most area of the sample. Position the left edge into the center of the live image field of view. 

 4. Using the ‘Tile Region Setup’ tools located directly below the image display, activate ‘Contour’. 

 5. Select the square contour tool 

 6. In the image display, click and drag to draw a yellow box around the live preview image. 

 7. Repeat step 5 and 6: using the XY joystick, move your stage position to the right, top, bottom most area of the sample. Position the top edge into the center of the live image field of view. 

 8. Once tile locations have been verified, close the Advanced Setup imaging window.                                              

 9. From the ‘Options’ menu in the Tiles dialog, set your Tile Overlap to between 10%- 20% to start. Note: The more detail you have in the overlap area, the lower you can set your tile overlap percentage. 

 10. Click ‘Start Experiment’ to begin tile acquisition. Note: For Brightfield Imaging – shading correction can be acquired from an area void of any sample. Defocus slightly to eliminate any dust or debris from the stage. It is also critical that white balance be accurate and done before tiling. 

 11. For tiling with AxioCam HR, select the orientation option 'flip vertically' from the dropdown menu in the 'model specific' tab under Acquisition parameters. 

 

 12. For tiling with AxioCam color, select the orientation option 'Rotate 180' from the dropdown menu in the 'model specific' tab under Acquisition parameters. 

 

   

 Shutting Down the System 

 1. Close Zen software 

 2. Transfer your data to the core server under your PI’s NETID: \\core-server.igb.illinois.edu 

 3. Logoff 

 4. Press Power button on EMS 3 box and wait for microscope to shutdown 

 5. Turn off X-Cite epi-fluorescence light source

Basic Optics
Initial Training for Optical Microscope Users

Data Storage
Store data on the data drive on the local machine: 

 D\Data may be deleted at anytime.

 Do not leave your data here!  

   

 The core staff will clean the computer drives occasionally. This means that your profile, desktop and data from the data drive will all be removed. We will also remove other files that look like they are not needed. 

 Core-Server: 

 The core-server was set up by the IGB Computer Network and Research Group (CNRG) as a place for core users to store their data long term and move it back to your office. You do not need to bring thumb drives or other devices that could bring a computer virus into the core when you come to collect data. Move your data off of the local machine onto the core server at the end of each imaging session. Your PI will have a folder on the core-server and you can make a sub folder for your work. you will have access to all of the sub folders in your PI's folder but not other PI's folders. All of the data in your PI's folder was paid for by your PI and belongs to him/her. 

 Your PI will be charged $8.75/terabyte every month  Email help@igb.illinois.edu   for information on charges and tape backup for long term storage.  

 Long Term Storage 

 CNRG provides tape backup for long term storage for $200/ terabyte 

  

Fluorescence Imaging
Why Fluorescence: 

 We can label what we want to see 

 

   

 Excitation and Emission 

 Dapi 

 

   

 

   

   

 

   

 

   

 Widefield vs Confocal 

   

 

   

   

 https://www.journals.uchicago.edu/doi/full/10.1086/689588 

   

   

  

IGB Core Instrument LSM 880
  

  

IGB Core Instrument LSM 900
  

 

 405nm, 488nm, 561nm, 640nm excitation. 

 Zen Blue 

 file:///C:/Users/gfried/Downloads/EN_poster_Beampath-LSM-900_A1.pdf

IGB Core Instrument V16


IGB Core Instruments Axiovert 200M
Axiovert 200M 

 

 Cameras 

 cMOS 

   

   

   

   

  

IGB Core Instruments LSM 700
LSM 700 

   

 

 http://zeiss-campus.magnet.fsu.edu/tutorials/spectralimaging/lsm700/indexflash.html

IGB Core Instruments Zeiss LSM 710
Light Path 

 

 https://www.gu.se/en/core-facilities/lsm-710-nlo 

 

 

 Seven visible excitation lines: 405nm, 458nm, 488nm, 514nm, 561nm, 594nm, 633nm. 

 Tisaphire laser 700nm to 980nm 

 Spectral Unmixing http://zeiss-campus.magnet.fsu.edu/articles/spectralimaging/introduction.html 

 Multiphoton Microscopy http://zeiss-campus.magnet.fsu.edu/referencelibrary/multiphoton.html  

 Fluorescence Lifetime Imaging Microscopy (FLIM) http://www.iss.com/microscopy/components/FastFLIM.html 

  

Objectives


Optics
Properties of light 

 Wave particle duality: 

 Light is a wave 

 

 

 Absorption and Emission 

 Beer's law 

 Refraction 

 Ray tracing 

   

 Resolution 

 Resolution: The ability to separate two objects 

 

   

 A definition of Numerical Aperture 

   

 

 Fourier transform: 

 Transform from real space to frequency space 

   

 

 Now we look at a real square wave and frequency space 

   

 

   

 Look at a simple optical system: 

 

 Mathematical prediction of the Point Spread Function (PSF) 

 on the left we have the mathematical point source know as a delta function. 

 

 were 

 

 the intensity at the Fourier plain can be found by taking the Fourier transform of this function. 

 or  

   

 

 this has the same intensity at all points inside the aperture and zero outside. The second lens is now taking a Fourier transform on a box function the width of the aperture. 

 

 or 

 

 Substituting in the definition for NA 

 

 

 

 

 Now we go back to Resolution. How close together we can position two points and still distinguish them 

 

   

   

 

   

 

 

 

 

 

   

 

 

   

 More intuitive approach 

 

   

   

 Notes from: 

 http://web.mit.edu/2.710/Fall06/2.710-wk12-b-sl.pdf 

 https://links.uwaterloo.ca/amath353docs/set11.pdf 

 https://www.thefouriertransform.com/pairs/box.php 

 http://www.phys.unm.edu/msbahae/Optics%20Lab/Fourier%20Optics.pdf 

 How to Chose the Optimal Objective Dr. Sebastian Gliem 

 Super Resolution Techniques 

  

Sampling
How does digital sampling affect resolution 

 Look at imaging these object with a digital camera 

 How close together do pixels need to be? 

 

 Sampling over time: 

 How often do you need to image a moving sample 

 

   

 Nyquist theory states that you should sample more than 2 X the frequency that you expect. 

 Over sampling 

 

 Nyquist sampling 

 

 Under sampling causes aliasing 

 

 Optical Transfer Function MTF 

   

   

   

   

 When objects get close together the contrast decreases. 

 MTF = Image Modulation/Object Modulation 

 MTF = 2(φ - cosφsinφ)/π   and

 

 

 φ = cos -1 (λν/2NA) 

   

 The Optical Transfer function is the Modulation transfer function times a phase component. 

 OTF = MTF × eiφ(f) 

   

   

   

 Camera bit depth 

 

 0 or 1 

 00 or 01 or 10 or 11 

 000 or 001 or 010 or 011 or 100 or 101 or 110 or 111 

 and so on 

 

 

 Jpg is 8 bit Tiff can be 16 bit 

   

 references 

 https://imb.uq.edu.au/research/facilities/microscopy/training-manuals/microscopy-online-resources/image-capture/nyquist-conditions 

 https://microscopy.berkeley.edu/courses/dib/sections/02images/sampling.html 

 https://ocw.mit.edu/courses/mechanical-engineering/2-71-optics-spring-2009/video-lectures/lecture-22-coherent-and-incoherent-imaging/MIT2_71S09_lec22.pdf 

  

Working in the IGB Core
Expect to walk into a room with a fully functional instrument 

 Let a core staff person know if you see a problem 

 Clean up when you leave 

 Acknowledge the IGB Core as: 

 

 “Core Facilities at the Carl R. Woese Institute for Genomic Biology" 

 Let us know when you publish 

 Collaborations with the core facilities staff can be beneficial in the development of unique methods or capabilities. 

  

Cryostat

video


EMPANADA Guide
To begin, I would recommend that one starts with the official empanada guidebooks as located here: https://empanada.readthedocs.io/en/latest/empanada-napari.html 

 Much of what I will be talking about is found here, but I will go over a few caveats that I've found in the process and try to explain the segmentation workflow in a step by step process. 

 Opening the software: 

 First, ensure that you have the software installed. This is best done by using the anaconda package manager. Through this, you can install NAPARI, the image viewer that EMPANADA runs. Once those are installed, you will have to activate the instance through the miniconda terminal by typing "napari". Note that if you made a virtual environment for running napari in, you will first have to run the command "conda activate [NAME OF INSTANCE]", with [NAME OF INSTANCE] replaced to what you named your virtual environment. Creating a virtual environment makes keeping track of dependencies easy, as NAPARI requires an older version of python than the machine may run. You should then be greeted by an interface that looks like this: 

 

 Loading a model: 

 By default, Emapanda comes with it's own model, MitoNet. As you can guess it is used for identifying mitochondria. The IGB core staff have also developed a primitive model for identifying the cores of myelin sheaths. You can tenatively find this model in the core server at \\core-server.igb.illinois.edu\groups\core\core server\shared\2023\empanada-tests\mylin_sheaths\tuned_data\mylin_test\unknown under the file name PanopticDeepLabPR_Mylinium.pth. With the empanada plugin loaded, go to the "Register a model" option, then follow this path and click on the proper .pth model to register it into the program. 

 

 Loading an image: 

 There are three methods of opening images in the napari image viewer. You can open them as a single image, a stack, or a folder. The image option will allow you to select a single image, whereas the stack and folder option will let you either select a certain subsection of the folder, or the entire folder. Be careful, napari will not verify your selections, so make sure you are only selecting images. 

 Running an image segmentation: 

 There are two options to run an instance of a segmentation that may seem similar, but do slightly different tasks. The 2D image segmentation program is simpler than its 3D counterpart. Each layer of image is taken into account separately than in the 3D version. This means that sometimes layers will be over segmented in the 2D version meaning that the program tries to give the same part of an image a label more than once. This typically does not happen in the 3D version. The tradeoff is that the 2D version is often faster than the 3D version, so it is advisable that one use it for basic parameter testing and other tasks, and then when you have verified your results, bring it into the 3D segmentation version. 

 Here are some of the options you will see and what they all mean: 

 

 NOTE: Raw images from the microscopes in the lab will often be 32bit TIFFs, these will not work in empanada! You must convert them into either 16 bit or 8 bit tiffs for segmentation to not throw an error! 

 image layer: the current image set you are working on. 

 model: which segmentation model you are running 

 use gpu: use the graphical processing unit (GPU) for faster processing (our machine are equipped with very good GPUs, so you will almost always want to run this) 

 use quantized model: enables better performance for cpu models,  be careful, this operation has precedence over GPU mode! 

 multi GPU: use multiple GPUs, our machines don't have them, so this option is meaningless 

 Image downsampling: If your image is oversampling, a trick could be this option, which basically scales down your image by a factor of the number you choose. This can be helpful for oversampling, but watch out for smaller segments getting lost. 

 Segmentation confidence thr(eshhold): Tells the computer how confident you'd like it to be in its selection of segments. Higher confidence means less segments, lower confidence means more, but better chances for false positives. 

 Center confidence thr(eshhold): This tells the computer how confident to be in where the center of the desired target is, it is not as important as the segmentation confidence, but can sometimes help prevent segmentation leaks if cells are tightly packed. 

 Fine boundaries: Much like the previous option, helps prevent segmentation leakage into other cells/organelles if your boundaries aren't well defined. 

 Semantic only: Only judges a segmentation by its shape, not any of its details, I generally wouldn't recommend this option. 

 (These next parameters are exclusive to the 3D version of the software) 

 Median Filter Size: the number of image layers to apply a median filter to, which smooths out the image in order to make semantic segmentation easier, but may distort tiny details that could be critical to detection. 

 Min Size (Voxels): The smallest object that is allowed to be segmented instead of filtered out as noise. 

 Min Box Extent: The size of a minimum bounding box to make sure there aren't an long spaghetti strand like segments that are often useless. 

 Max objects per class (also found in 2D): maximum segmentations per image classifier if a model segments more than one type. 

 Inference plane: which plane to go through as your microscope did to get a proper 3D stack, xy is often sufficient. 

 Run ortho plane: This is the function that turns your AI segmentation into 3D models.  NOTE: This does not work with our current microscopes as almost all of them are anisotropic! 

 Return xy, xz, yz stacks: Return every stack that the software generates in the 3 possible planes. 

 Voxel vote thr out of 3: this is the amount of layers that a segmentation must be present in to be displayed in the ortho plane. 

 Note that the tick box "permit detections found in 1 stack consensus" is the equivalent of setting the top option to 1. 

 Zarr storage is useful if your machine doesn't have enough ram to work with your image sets (typically workstation machines have anywhere from 8-256, if your image set is bigger than this, you'll likely want to use it), it stores the segmentations temporarily on disk. 

 Directory: the directory where these temporary stacks will be stored 

 Chunk size: how big you want your temporary images to be, as they stay saved on disk and can be extracted if desired. 

 Training a model: 

 As a researcher or core staff assistant, you will find situations other than myelin sheaths and mitochondria that you or others will want segmented. This will require two key components. The first is training data, and the second is segmented data corresponding to the desired organelle of the training data. Once you have these two things, you can begin training a model. Empanada requires a certain directory layout for the data, you can achieve this in an easy manner by using the store dataset tool. Just specify the images and the segmentations and which directory you'd like to output this to. 

 Next thing is to do is to run the "train model" module, which looks like this: 

 

 The options for this are as follows: 

 Model name: the name of the model you wish to create, make sure it is descriptive of the organelle you are targeting to avoid confusion. 

 Train directory: The directory of the unsegmented images, NOTE that this should be the top level above the folder which contains the folders for the properly formatted images and segments if done with the store database command. 

 Validation directory: The directory in which the segmentations are contains. This says it is optional, but note that the segmentations are not optional in themselves, but rather this option should be used if the segmentation directory lives in a separate directory from the images, if no input is made, the program will use the same directory from the train directory. 

 Dataset labels: When segmenting your data, you will have be able to mark different types of organelles in your cell. These should be marked with labels that fit into a certain section of label numbers. For instance, you could mark mitochondria in the 1000s, golgi complex in the 2000s and so on. How you define these ranges is up to you, but make sure you mark the boundary in the label divisor option. In this option, you will have to specify each group of segmented organelle (this can be a simple list of 1 through the amount of groups, this is just to tell the computer which groups are different), the name, and segmentation type, semantic or instance. Semantic segmentation is solely based on the shape of the object, but instance segmentation is based on the shape, placement, and details, and as such is generally more useful. 

 Here is an example of a proper format: 

 1,golgi,semantic 

 2,ER,instance 

 3,mitochondira,instance 

 Model architecture: Which architecture you want to use for training your model. 

 Fine tunable layers: Denotes which stages in the architecture to apply fine tuning to. This should generally be set to "All". 

 Iterations: How many times the model should be trained on the dataset. The default 500 generally gives the best results, but if you want to shorten the amount of training time (this could easily be tens of hours), then reducing the iterations will help. 

 Patch size in pixels: The general size of segmentations in pixels, must be divisible by 16. 

 Custom config: You can use this to specify a config file with all the options pre-set. May be useful if you are trying to train many similar models for comparisons. 

 Description: A free text box in which you can tell any future users any details about your model, such as what it was trained on, best use parameters, and possible weaknesses of your model. 

 Once this is all set up, you should have to wait somewhere between 10 and 20+ hours depending on how big your image set is. After that, you should see a .pth file appear in wherever you specified your target directory to be. If your model did not automatically load into the software, you can use the register a model program mentioned previously. 

 Finetuning a model: 

 Finetuning a model is normally meant to clean up segmentation rather than add new data to your model. You can use this if you want to simply refine your model, as it will take much less time to finetune data you already have versus add new data and regenerate your model. Normally the workflow for this is as follows: Generate segmentation on a sample. Clean up any errors in the segmentation. Use the "pick patches" tool to select 16 or more (software limitation) segmentations to feed the computer. Store them as a database. Run the database through the finetuning tool. The finetuner has almost all of the same options as the trainer, so I will not repeat my word in this section, simply check the train a model documentation. 

 Other tools: 

 The software suite consists of a number of other tools that are for ease of use. These are not critical to the training or segmentation process, so I will only cover these tools briefly. 

 Merge labels: 

 This tool merges two of the layers on the left hand side of the screen (don't get this confused with merging segments, which can simply be done with the paint bucket tool). To use this, you will need to go to your stack of choice. From there you will mark the layers that you want merged with the point tool. 

 Delete labels: 

 Much like the merge labels tools, this one functions quite similarly, select the stack you'd like to delete from, use the point layers to select which layers from the stack should be subject to deletion. 

 Split labels: 

 This will put any segmentations within a certain range of the point placed on a separate layer from the currently selected layer, you will need your points layer and your stack as arguments once again. You can also adjust a slider to tell the computer what radius from the point should be considered. 

 Jump to label: 

 Given a label ID, jump directly to it. Though it may be easier to simply use the in built NAPARI arrow buttons at the top left to simply cycle through which label you want to interact with.

Köhler  Illumination
Köhler Illumination for Brightfield 

 Köhler illumination is a method for evenly illuminating the sample with white light. This method is very important for brightfield imaging in widefield microscopy, and for using the T-PMT for confocal imaging. Brightfield illumination from light source focuses onto the sample via condenser, as a result extra steps are necessary to ensure the light-path is focused on the sample. 

 Bring the sample into focus 

 Close the field diaphragm ( 2 ) until you can see at least one edge 

 Adjust condenser height ( 3 ) until the edges of the diaphragm are crisp. You’ll see an octagon. 

 Center the diaphragm ( 4 ) mage using the two centering screws 

 Open the field diaphragm (2) just until the image fills the field of view (FOV) 

 

 Link for setting up Köhler illumination 

 ZEISS Microscopy How-to: Set up Köhler Illumination on your ZEISS Axio Imager (Upright): 

 https://www.youtube.com/watch?v=Qk8dbT-N1xw  

 MicroscopyU has a great interactive on guide for Köhler illumination:  https://www.microscopyu.com/tutorials/kohler 

   

   

  

Lsm 700 Info

LSM 700 Basic Operation
  

 Startup 

 Turn on the  laser lamp 

 

 

 

 Login into the  computer 

 Start  [ZEN]  software - Start System 

   

 Visual Operation 

   

 Loading a slide  (#1.5 cover slides are the best) 

 Set the objective to  10x  using the touch pad on the microscope (recommended to always focus first on 10x – as the objectives are parfocal you can then go up in magnification as necessary while remaining in focus). 

 Click on load button on knob attached to microscope  [Microsope] [Load position] 

   

 

 

 

 

 Tilt transmitted light source back by placing one hand on the back side of the base of the eyepieces and one hand on the white part of the transmitted light source.

 

 

 Load the slide into the slide holder cover slip down (to move the sliders apart press down on both buttons slide as necessary) 

 Click  on working position knob [behind load position]  to return to working position –  Be careful to make sure the slide does not run into the objective  (this is why you should always focus on 10x first) 

 Note: Do not depend on the preset load and work settings. Samples and slides of different thicknesses may have been used by the previous investigator. 

 Tilt the transmitted light source back into place 

 Focus on your sample by using either transmitted light or fluorescence:

 This can be done in a number of ways: 

 §  Method of choice  - In Zen Select  [Locate] 

 Select  [Online] 

 Select the desired fluorophore or transmitted light 

 Locate region of interest and focus the samples 

 Select  [Light Off]  to prevent bleaching § Using the touch pad on the microscope §§ Using the buttons on the microscope itself and touchpad. Choose TR (transmitted light) or RL (reflected light and the filter: DAPI/ 488 /Rhod). 

 Focus the specimen. With coaxial knob you can move the sample on the x-y level. 

 

 Change the objective as necessary for your imaging. 

 To add oil (only for the 40x and the 63x) – remove the slide 

 

 Open the lid of the bottle and scoop some of the oil with the plastic oil probe. 

 Turn it sideways (so that the longest edge is vertical) – there should not be oil dripping off of it 

 Move it over the objective and turn it flat – one drop should fall onto the center of the objective 

 Never touch the oil probe to the objective 

 If you need to return to a lower magnification from an oil objective you must use the 10x.  (The 20x has a short working distance and will get oil on it, which is bad for the objective and will hinder imaging with it. Additionally, the 20x objective is a high NA objective and is therefore expensive) 

   

 Imaging 

 Select  [Acquisition] 

 

 

 Load a user profile image file Select  [Reuse]  or use  [Smart Setup] 

 Select  [Dye] , select show all, click on arrow and pick dyes (do this for each fluorophore your sample is labeled with). This is where you pick the pseudocolor for each channel. 

 o Pick imaging sequence: 

 § Fastest: 2 channels per track (meaning 2 channels are imaged at the same time with 2 PMTs) 

 § Best = slowest: each channel is in its own track and is imaged separately (this prevents bleed through) 

 § Best-compromise = some channels are in a track together and others are in their own tracks (medium time to get image) 

 Select  [light path]:  select track (if not showing click show all manual settings towards the top left of the software window)

 If possible it is best move the VSD (variable secondary dichroic) to be at the same nm for all the tracks 

 § VSD is the white vertical line on the wave length graph and the nm is listed under Split 

 

 You can use filters (show all and click on the arrow in the center of the colored bar below each emission spectra to see the filter options) to further narrow the emission collection 

 For  acquisition  o Settings will depend on your sample. A good starting point is:

 Scan mode: Frame o Frame size: 1024 x 1024 o Line step: 1 (always!) 

 Speed: 7-9 o Average: 1-4 o Mode: line 

 Method: mean o Bit depth: 8 -16 bit 

 Unidirectional scanning (arrow points in only one direction) or Bidirectional scanning makes imaging faster. 

 

 For  scan  o Select one track by unclicking the other tracks o Live: center, orient and focus samples (use one track) o Turn on all the tracks

 In the  [Channels]  menu adjust the  [laser power],  2% default (1-5% is usually all you will need), and  [pinhole,  1 airy unit – 1 AU ] 

 

 

 

 

 

 

 Select  [Live] 

 Select  [Dimensions]:  (sometimes this is easier to do with one channel or track selected at a time) 

 § use  range indicator mode  (click on color of channel and it will automatically switch to this or by clicking the color field in the View-Dimensions you get the scanned image in a false-color. Red color indicates saturated pixels (maximum) and Blue = zero (minimum). To avoid saturated pixels set the laser power and gain so that the red pixels just disappear 

 § split channels 

 § Select one channel within a track by clicking on it (light gray bar line is the selected one)

 Focus on the brightest plane of this channel in your sample 

 adjust  [Gain] , to just below when red (overexposed) pixels appear (you want this between 500-750; if it is too low then lower the laser power) o Gain = voltage on PMT. Higher number than you can detect weaker signals and also more noise. 

 

 

 o On histogram you want to shoot for 75% of the range 

 Adjust  [offset]  – only move up or down by 1-2 to get background to blue speckles (starry night) 

 [Digital gain]  set to 1.0, do not change. 

 Repeat for all the channels and all the tracks 

 Click  STOP  –for stopping the current scan procedure 

 To take a single image: Select  [Snap] to collect the final image and add a scale bar 

 

 

 

 § Save image to groups-server or to drive D: user file deleted anytime folder 

 

 

 

 § Always save as a lsm/czi file first before exporting as tiff. In CZI-format all the information and hardware settings are saved together with the image. Export -function saves the contents of the image display window including analysis and overlays in selected format 

 

 

 

 You may also export a tiff file 

 

 

 

 

 

 

 §  Do not store on drive C:\ 

 § Once you have moved your data to the core server please delete it on the confocal computer (any images over one month old will automatically be deleted) 

 Z-stack (select at upper left) 

 

 

 

 

 

 

 § Choose one track to focus 

 § To optimize interval click [optimal] 

 § Set the first and last 

 

 

 

 

 

 

 

 

 

 

 

 

 § Re-select all the tracks 

 

 

 

 § Start experiment (upper left window) 

 

 

 

 § If you have a dramatic loss of brightness as you reach the end of your stack see core staff about Auto Z Brightness Correction 

 Tile Scan (select at upper left) 

 

 

 

 Click on show all and select tile (example 5x5 for 25 images) or (3 x 3) 

 

 

 

 

 

 

 

 

 

 

 

 

 § Alternatively, for 10 or 20x just select the size of the image

 Centered = current image is the very center of the size you select 

 Bounding grid lets you scan and move to positions an dmark 

 Convex hull generates an image the goes in a line from point a to point b. 

 Do not try to crop a tiled imaged by the crop tool – change the size using the image size (um) at the top of the right column. 

 If you see image issues, then set overlap to 10%. 

 § If you are using the 40 or 63x and doing tiling set the overlap to 10% 

 § Start experiment (upper left window) 

 You may export tiff file 

 

 

 

 

 o  Position (select at upper left) 

 § Mark positions – marks X, Y, and Z. 

 If you want to do Z stacks at multiple positions, then each Z stack will be the same number of slices. Therefore, you need to make the Z position setting to be in the middle of your sample and then set the Z stack to go so many microns above and below your setting. 

 § Autofocus option 

 Fl = fluorescence is used to find sample. Takes a stack before it images and uses the brightest plane as the center. This can cause sample bleaching issues 

 Reflection = uses the transmitted light to detect the coverslip and you set the focus to be a certain number of microns above the converslip. 

 If you sample is sloshing around in liquid – none of these autofocus options will really help 

 

 Bleaching = photomanipulation (select at upper left) 

 § Time series and regions windows open by default 

 § Scan image 

 § Select the  [Regions]  and click show all:

 Draw desired shape or shapes 

 Do not check acquisition box or you will only image the bleached region 

 The scope will scan the whole way across the X direction and only bleach the region boxed. This means your box should always be longest in the X direction (you can always rotate your sample). 

 Zoom Bleach – doesn’t scan all the way across in the X direction so it is faster, but less accurate. 

 §  [Bleaching]  and click show all: 

 Check start bleaching after a number of scans (2 is good) 

 Set - Repeat bleach after # of scans (pick a number) 

 Set - Stop when intensity drops to % starting intensity (after this it will no longer bleach, but it will continue to image) 

 Iterations  – 20-100 is a good place to start (this and laser power are the points to adjust) 

 Select laser to use to bleach and laser power ( 25%) 

 §  [Time Series]  and click show all 

 Select number of cycles = number of time points. 

 Leave interval at 0 – usually you want to see this as fast as possible § Notes: 

 In the gallery view, the green boxes mark the upper left corner of the image that was taken before the bleach. 

 We currently do not have the analysis plugin for FRAP – you can analyze the data in Image J or metamorph. 

 Please note that when you select a region it will scan across the full X and only bleach in the region of interest. So if you have an oblong or rectangle region you want to bleach it is fastest to orient the longest part along the X plane. 

 

   

 When your session is done, make sure: 

 You have transferred your files to IGB group server / lab folder 

 

 

 

 

 Clean oil off 40x and 63x objectives with lens paper not kim wipes. 

 Return to 10x objective 

 Follow shutdown procedure 

   

 Shutdown   

 Check calendar to determine if someone is scheduled after you (within 1 hour). 

 

 

 IF Yes: 

 Turn off the software and logout 

 

 

 IF No: 

 Exit the Zen software and log off computer 

 

 

 

 Turn off laser lamp 

 Don’t turn of the microscope or anything else. 

 

 If you have any problems: 

 Always report any problems to Core Staff 

   

 Viewing and Processing Data 

 To view and process you images, ZEN 2012 lite freeware is available in  www.zeiss.com/lsm . Fill the form and you will receive a link for installation via email

LSM 710

FCS module


FLIM module


LSM 710 Basic Operation
Turning On the System 

 Make sure the main power switch, systems switch and components switch are all on. The key should be turned on, please do not touch the key. 

 

 Turn on the X-Cite lamp next to the microscope. Turn on the Argon ion laser. Turn the key to the horizontal position and flip the small metal switch up. If the light is red lower the current control knob until it is green. 

 

 Loading Your Sample 

 To load your sample push the arm of the microscope back until it holds in place (don't grab it by the condenser).  Before loading the sample, press the lower button on the focus knob to lower the objective to the load position. 

 

 Move the sample to holder to the correct position for either a slide or a 35mm dish.  Make sure to load the sample with the coverslip facing down (towards the objective).  If you need to use the 40x or 63x objective add a drop of water or immersion oil to the objective lens. After placing the sample on the holder, press the upper button on the focus know to return the objective to the working position.  Lower the microscope arm (there is an interlock that will prevent the laser from turning on if the arm is in the raised position). 

 

 Locating Your Sample 

 On the computer, login with your IGB account and password, and open the Zen program.  Select "Start System" from the options it presents you. Wait for Zen to load all the components and connect to the microscope. Zen will present you with four tabs at the top: Locate, Acquisition, Processing, and Maintain.  It will start you in the Locate tab; this will allow you to find the sample in the eyepiece. To find the sample in bright field, turn on the lamp, transmitted light (TL) shutter as shown below in the image on the left.  You will also need to choose the empty filter from the filter turret. After those items are selected, You can look in the eyepiece and adjust the course or fine focus know to find your sample.  To find your sample with the 488 or 561 filter, close the TL shutter and switch to the green or red filter as shown in the picture on the right.  If you don't see blue or green light coming from the the objective, check that the manual shutter on the X-cite lamp is open.  Use the focus knob to find your sample in the eyepiece. 

 

 You can also use the touch screen to control these settings and view the sample in the eyepiece. The main menu on the touch screen will display information about the objective and stage position.  There is also a button labeled "Make It Visible" that will put the microscope directly into a bright field configuration so you can view the sample. 

 

   

   

   

   

   

   

   

   

   

   

 If you press the microscope button you'll be able to select the objective and reflector.  Once you have found your sample in the eyepiece, you can move on to setting up your scan. 

 Setting Up Your Scan 

 Once you have located your sample in the eyepiece, click the acquisition tab (not the menu item at the top).  This is where you will setup the parameters for your image. Zen will show you a series of menus, the four most relevant (Lasers, Imaging Setup, Acquisition Mode, and Channels) will be open and the rest will be collapsed.  We will go through the four menus needed to setup a scan.  Also, we will be using the live and snap buttons at the top of the screen as shown below. 

 

 In the laser menu, turn on the 561 laser is you need it.  The laser properties menu will show you the status of the laser, it should say either "connected" or "warming up".  If it says "inaccessible" please contact the core staff. 

 

 Once the lasers are on.  You can proceed to the "Imaging Setup" menu.  Press the "+" button to add a track, you'll want to add a channel for each dye in your sample.  You can have up to four tracks.  I'd also recommend changing the "switch track" option from line to frame.  Switching the track every line requires that all of the optics in the light path be fixed, if you aren't careful this can be frustrating. 

 

 For each track choose a laser.  Then choose a beam splitter (dichroic filter) that matches the laser line. For example, if you are using  the 488nm laser, choose one of the beam splitters labeled for 488nm.

 

 

 Next, select the detector you want to use.  By default channel 1 is selected as the detector.  As long as  "switch track every frame" is selected you can just leave the detector as channel 1.  Choose a dye from the database, this will display the emission spectrum of the dye.  Adjust the emission range so that you are collecting most of the light from the emission spectrum.    Repeat this for every track - one track for every dye in the sample.

 

 Once the tracks are all setup, move on to the channels menu. At this point it is worth noting that "tracks" refer to the light path setup and "channels" refer to detectors.  There are cases where you might want to do more than one channel per track, but for now we will just focus on using on channel for every track.  

 One important user interface note, the check mark box can be unchecked and the the computer will ignore that track.  This is different than highlighting a track/channel which will display the parameters for that channel. 

 

 For each channel you will need to adjust the laser power, gain, and pinhole size.  Start by setting the gain to 700.  Raising the gain will increase your signal but will also increase your noise.  Lowering the gain will lower the signal and noise, requiring you to use a higher laser power.  We have found that 700 gain is a good starting point.

 

 For the pinhole, start with the middle channel (or the most important channel).  Press the 1AU button.  This will set the pinhole to one airy unit (the diameter of the airy disk).  Next to 1AU or 0.99 AU is an approximation of the optical section thickness, 1.5 microns in this case.  The section thickness will increase as you open the pinhole.  It is important to keep the pinhole diameter (26.3 in this example) the same across all channels.

 

 Now you need to adjust the laser power (just above the pinhole).  It starts at 0.20% and can be raised to 100%.  You will need visual feedback to correctly adjust this.  Uncheck all but one track, and click on the the "live" live button at the top of the screen. At first you might not see anything, it is very likely the sample is not exactly in focus. Click on the "range indicator" button at the bottom of the screen.  This button will turn off the false color and display the live image in a diagnostic color scheme.  Blue is zero, black to white is signal, and red is saturation.  

 

 You can see in this image that  there is signal, but it is weak.  First adjust focus until the signal gets brighter (black to grey to white to red)..  

 

 Maximize the amount of red to make sure your sample is in focus.  Then lower the laser power to that only a few red pixels are visible.  You want to use as much of the detector's dynamic range as possible without saturating the detector.   A few red pixels are okay, you just don't want big patches of them. The correctly adjusted image should look as shown below. 

 

 Press the "stop" button, and repeat this procedure for the other channels.  Once you are done setting up all the channels we will setup the scan parameters in the acquisition menu 

 

 Start with the scan area, you can adjust the size, position, and orientation of the scan area.  Adjust the size and position to frame the sample.  There are two important numbers displayed next to the scan area.  The "Image Size" is physical size of the image, and the pixel size is the image size divided by the the frame size.   Click on the XY button to choose a frame size.  If the frame size is too small, then the pixel size will be too big and your resolution will suffer.  Click on the "optimal" button just below the XY button and Zen will select the correct frame size.

 

 You'll need to choose a scan speed.  This will affect the scan time and pixel dwell time.  The pixel dwell time is how long the detector collects light at each pixel.  I recommend not going below 1 microsecond, as this will result in a noisier image.  You can choose an averaging number to average lines/frames together in order to improve the signal to noise ratio.  In general keep the bit depth at 8-bit.  Once the settings are correct select the "Snap" button to start the scan.  One note, "snap" will use all the settings in the acquisition menu.  The "Live" button with always use a 512 x 512 frame size and scan as fast as possible.

 

 

 Once you have taken the image, save it as a CZI file on the D drive in the "User Files Deleted Anytime" folder.  We don't guarantee files saved here are kept forever but we don't go out of our way to delete them.  

 The next time you use the LSM 710, you can open this CZI file and click the "Reuse" button just below the range indicator button, this will reapply all the settings used to take the image. You'll still want to check the laser power settings with the range indicator as this will change from sample to sample.   

LSM 900


MINFLUX
Molecular resolution in 2D and 3D

Introduction
Minflux is a florescence imaging and tracking technique using a 2D doughnut or 3D bottle beam to localize switchable fluorophores to single digit nanometer resolution. 

 Expanding on this 

 Minflux turns on fluorophores using a 405 laser. This allows the microscopist to control the number of active fluorophores and eliminates the situation where two active fluorophores are close together. 

 Minflux scans a sample with a doughnut shaped beam when an activated fluorophore is found (red star) in a field of fluorophores in a dark state (circles) the center of the excitation doughnut (green spot) moves in a hexagonal pattern 2D or octahedral pattern in 3D with a spherical excitation pattern, to determine the location of the fluorophore. The search pattern is minimized sequential. 

 

 From Minflux unrivaled resolution Abberior 

 Light Path 

 

 Sample preparation 

 Cells are seeded on 18mm glass coverslips (No 1.5 or 1.5H); or chambered coverslips (no plastics cover slips). Fixation is done as usual (Formaldehyde or Methanol). 

 Minflux uses labeling techniques developed for dSTORM, so this means that Alexa fluor 647 is common. along with an oxygen scavenging GLOX buffer.  β-mercaptoethylamine (MEA) is used as a blinking agent. Fiducials are also used. 

 Sample mounting and imaging buffers. For the sample stabilization during MINFLUX measurements, gold nanorods (A12-40-980-CTAB-DIH-1-25, Nanopartz Inc.) are used as fiducials. In brief, an undiluted dispersion of the nanorods is applied to the ready-made samples. Before mounting the samples in imaging buffer, the coverslips are rinsed several times with phosphate buffered saline (PBS) to remove unbound nanorods. For MINFLUX imaging of samples labeled with Alexa Fluor 647, GLOX buffer (50mM TRIS/HCl, 10mM NaCl, 10% (w/v) Glucose, pH 8.0, 64 μg/ml catalase, 0.4 mg/ml glucose oxidase, 10–25mM MEA) is used. Samples are sealed with twinsil (picodent).  Ref 1 

 Labeling Dyes 

 Live Cell Imaging (PALM) and Single Molecule Tracking:  

 - Photo-switchable Organic Dyes/ Proteins, e.g., Janelia Fluor 549 or 646 dyes (Tocris) - Photoconvertible Oganic Dyes/ Proteins, e.g., mMaple (plasmids sold by Allele Biotechnology) Fixed Cell Imaging (dSTORM): 

 -Carbocyanine dyes (sCy, ATTO, Alexa Fluor or Flux dyes may be used). Various dyes excited at 640 nm, such as Alexa Fluor 647, CF660C, CF680 or sCy5 can be used. Best Dye: Alexa 647 or Flux 647 for one color and sCy5 + CF680 or Flux 640 + Flux 680 (recommended) for 2 color-imaging.  

 Intracellular/Surface Live Cell Imaging (dSTORM): - SNAP, CLIP or Halo-tags for single color live cell imaging  - SNAP & CLIP-tags for 2-color - SNAP & Halo-tags for 2-color (preferred if already in cell line) - Nanobodies (Proteintech) 

 No DAPI or Hoechst staining in sample.  Labeling density should be very low (dilute antibodies 1:500 to 1:1000) to avoid over-labeling. 

 Background labelling: there should be no background. - increase blocking steps & times - increase washing steps & times - reduce antibody concentrations 

 Stabilization 

 Sub nanometer stabilization is required to measure the location of a fluorophore to nm resolution. This system preforms to 0.39nm in x and y and 0.6nm in Z. This performance is impressive since 980nm light is used. 

 980nm light (easily passes through tissue) passes through a half wave plate to rotate polarization and then through a polarizing beam splitter and relay lens to focus on the back aperture of the lens. A quarter wave plate changes the linearly polarized light to right-handed circularly polarized light. Reflected left-handed circularly polarized light changes to vertically polarized light and passes through the polarizing beam splitter, a spacial filter and travels to the camera. The spacial filter removes low spacial frequency information from the image. 

 

 figure from Ref 1 

 

 from supplement 

 gold nanorods rotating force: https://www.osapublishing.org/oe/fulltext.cfm?uri=oe-22-21-26005&id=303075 

 References 

 1 Schmidt R., et al. (2021) MINFLUX nanometer-scale 3D imaging and microsecond-range tracking on a common fluorescence microscope. Nat Commun., 12(1):1478. 

 2 Balzarotti, F. , et al. (2017). Nanometer resolution imaging and tracking of fluorescent molecules with minimal photon fluxes. Science, 355(6325), 606–612. 

 

 

 

  

Separating two signal based on dcr (detector channel ratio) in paraview
Goal: 

 to separate signal form 2 detectors (CY5 near vs CY5 far), especially when you have two dyes for example AF640 and AF680 

 1. Open image in Imspector 

 2. Minflux Data Panel → Data source: it is usually None , so you must select data source that you want to view in Paraview. 

 3. Minflux Data Panel → Launch Paraview                                      

 

 4. in Paraview → under localization tab →select coloring to dcr (detector channel ratio)    

 

 5. in Paraview → under localization tab → click right add filter → data analysis →histogram → select input array dcr → change bin count to 100 → press apply                                                                                                                                                                          

 

 From histogram plot we can see two peaks from CY near and CY far detector, we can separate the two channels for example here we want to discard dcr value from 0.32 to 0.36                              

 

 localizations tab: you will see color map editor or rainbow icon (edit color map) to show the color map editor          

 

 

 Choose Preset to change color map you want to use for example cold and hot and click apply          

 9. drag the color map (A) on the left side up (B) → we want to discard dcr value from 0.32 to 0.36 we will set this to black color: set the left circle the lowest dcr value 0.32 (enter the value on Data and click enter) and the right circle to 0.36 → make another two circles 0.32 and 0.36 on the middle circle double click on the center of circle then select color: choose black, repeat for the left circle for example select cyan HTML#00e9e9, and for the right circle select orange HTML #ff8206                                                                                                    

 10. Signal with dcr value from 0.14 to 0.32 in cyan and dcr value of from 0.36 to 0.7 in orange                                

 

                                                                                                                    

Superimpose images in Imspector
Goal: 

 Superimpose confocal images 

 Superimpose confocal image with Minflux image 

 Note: Images can be combined if they have the same length and the same pixel size. In general, the pixel size of the Minflux image usually is much bigger than confocal image. As a result, we need to do image interpolation to make sure the pixel size of the two images is the same. Length and pixel size of an image can be checked using change Stack Size by pressing CTRL T . msr file can be opened in image J. 

 Step by Step Superimpose images: 

 Example: combine a confocal image and a minflux image, both images have the same length however the pixel size are different.                                                                              

 Select an image and press Ctrl T 

 Change the pixel size of a confocal image to match the minflux image: go to Analysis and Interpolation , Change the resolution of confocal image to pixel size of minflux image, choose result format to float and press Now, we have similar length and pixel images.                                                                

 

 Choose new window                                                                                        

 Copy the image (Ctrl C or press A ) and paste (ctrl V or press B ) into the new window. When you paste the second image, warning appear “pixel dimension mismatch ignore and paste?”, select yes                                                                                                                                                                                                                                          

 Change the color map or each image or choose RGBize then superimpose the image using gallery mode or Ctrl G 

 An example superimposes confocal image with tracking image: 

   

                                                                                    

Tracking in ParaView
Goal: how to analysis tracking data in paraview 

 1 Imspector : open your image in Imspector 

 

 2 Minflux Data panel → select data set on Data source panel → Compile trace data (Trace data compiled on the bottom of Minflux Data panel) → then Launch Paraview                                                                                                            

 3 ParaView : 

 3.1 Localizations: show the dots where molecules were localized: 

 loc: localization coordinate; in raw data: determined position of the respective localization in x, y and z 

 cfr: center frequency ratio; emission frequency measured at the center pattern position (= efc) divided by emission frequency measured at pattern positions on the circle (= efo) 

 dcr: detection channel ratio; emission frequency measured on detector 1 divided by emission frequencies measured on detector 1 + detector 2 

 fbg: fluorescence background; determined background level (as count frequency) in the sample 

 efo: effective frequency at offset tcp position; emission frequency measured at the pattern positions on the circle (= ”offset”; not in the center position) 

 tid: trace ID/identifier 

 tim: time; timepoint of localization acquisition in respect to start of the measurement (in seconds) 

 itr: iteration 

 3.2 ExtracBlock1”: the connection between localizations 

 Hover Cells on (green triangle): if you click one of the traces this will show you ( block, trace, ID, type, spd, dst) 

 4 Traces → Histogram → select input Array (dst, dur, gri, len, mst, siz, tid, tim) 

 Dst: distance from the start point to the end point (it does not follow all the traces)                                                                  

 Dur: duration is the total time of the track in second                                                                                                      

 Gri: Grid is scanning point in iteration 0 (this not necessary for histogram) 

 Len is the length of completed trace (this is different from dst)                                                        

 mst????                                                                                        

 Size is the number of localizations in the trace                                                                                  

 5 Traces → ExtraBlock1 

 Loc: Location of each track, you can color coding based on X, Y or Z 

 Tid: random color coding of the traces, by using the traces ID, this similar with Imspector                                  

 Tim: each localization on the track is color coded regarding the time start of the trace to the end of the trace, (blue to red color coding means blue is the start of the trace and red is the end of the trace). Note: the first localization to the second seem longer than other track                                                                                                                                                                                                  

 Dst: distance from the start to the end point. Below by using Linear-Green color map show that dark green is the shortest and the white is the longest distance from start localization to the end point/localization.                                        

 

 Dst: length between two localizations, the figure we choose one trace dark green is the shortest distance and the white is the longest distance.                                                                                                                            

 

 Spd: distance between 2 localizations divided by time                                                                                         

Nanozoomer

Nanozoomer Basic Operation
Warning 

 Before beginning, have one of the core staff ( listed here ) look at the slides to confirm that there are no protruding coverslip edges.  The Nanozoomer is very particular about slide size, any non-standard slides or protruding coverslips will cause it to fail.  Also, make sure the slides have dried for 24 hours as any excess, wet mounting media can cause the scan to fail.  If the coverslip if protruding, you can use the sand paper near the Nanozoomer to file it down.  If a slide drops in the Nanozoomer, contact the core staff. 

 Loading Your Slides 

 Make sure the Nanozoomer is on. 

 Load the slides into the cassettes with the coverslip facing up. Push the Nanozoomer door open and place the cassette into the first slot.  You can put the 30 slides in each cassette and there are 7 cassettes, for 210 slides total. Note, if the yellow (busy) light or red (error) light are on then the door will be locked. 

 

 After the slides are loaded, close the door and log into the computer. Open the NDP Scan software. 

 Running a Scan 

 First we need to check the settings.  These settings are profile specific.  However, most users will use the same "Brightfield" profile. If you use the Nanozoomer often, please ask the core staff to make a specific profile for you.  We generally done encourage individual profiles because it eventually slows the software down. 

 Choose the "Settings" button 

 

 Change the file path you want to save to. Click apply to save changes. 

 

 Open the general settings 

 

 Leave the multiple scan area settings checked.  Make sure the "Brightfield" profile is selected and choose "Edit." 

 

 The "Region" settings will be displayed first.  You can leave the Scan Area as Automatic, the Focusing as 5pt Auto, and the thresholds at 10-90%. It is likely you want the coverslip filter enabled.  Choose a minimum tissue size so that the program will not identify small objects.  Set the Split Tissue setting to about 10mm.  Set Focus Pieces separately and excluded the Excluded Area (this will make scans faster).  

 If you plan to manually identify your tissue, these settings are less important.  If you plan to run the Nanozoomer in automatic mode, then it is important that these settings are correct.  You might want to click the the load next slide button.  This will load a slide and allow you to use the preview button to see how well the software is identifying the tissue region.  If the preview does not look good, adjust the thresholds and minimum size and try again. 

 Once you are done here click "Apply"" and then click "Output" at the top. 

 

 Just make sure the output path here is blank.  If someone has added a file path here, then it will over write the file path you added in the general menu.  Click "Apply" and then "OK" to return to the main menu. 

 Running a Batch of Slides 

 At the main menu, click "Batch of Slides" 

 

 At the top of the next screen, you'll need to choose your profile that we edited in the last section.  Unless we created a profile for you, this will be the Brightfield profile.  Choose Automatic or Semi-Automatic batch type.  For Automatic, just click "Start Batch" and let the scan run.  For Semi-Automatic, we need to verify the scan areas that the software suggests. Choose "Start Batch" to begin this process. 

 

 In Semi-Automatic mode, the Nanozoomer will load the first slide, take a quick black and white overview picture for the purposes of finding the tissue region, and then load the next slide.  It will take all of the overview pictures before moving on to scan the slides.

 

 

 You can right click on the the small image to see a bigger overview image.  To change selected area, click on the "Setup Scan" on the right of the screen. 

 

 If you don't like the regions that the computer selected, you can click on the "Scan Area" icon and choose to delete the scan areas. 

 

 You can use the left mouse button to create a yellow region that covers the tissue.  The blue rectangle cannot be adjusted directly, it is the boundary of all the yellow regions.  You can add as many yellow rectangles as you like, generally you want to have on rectangle per tissue.  You can use the right mouse button add focus points to the tissue.  Add about 5 focus points per tissue region.  Once you are happy with the scan regions, click "Accept" and the Nanozoomer will present the next overview image to you.  You can setup the scan regions as the Nanozoomer begins scanning slides.  If you don't see any issues with the scan region of the next few overview images, you can "Accept All."  This will start the scan for all slides. 

   

 

 As the scans begin, a progress bar will appear with an estimate of the time remaining. If focusing fails, this will also be displayed here.  Once a slide is scanned, the "Abort Scan" button will turn into a "View Scan" button, click this to open the slide in the NDPView2 software. 

 NDPView2 

 

 In NDPView2 ( downloadable here ) you can zoom in and move around in the slide you just scanned.  If you move your mouse to the left of the screen, You can add a scale bar, annotations, and make color corrections to the slide.  Moving the mouse all the way to the right will give you an option to select a zoom level (1.25x to 40x). 

 To save a tiff image, you can right click and choose to export as tiff.  Be careful doing this, by default the image will only show the current field of view at the computer monitor's resolution (3840 x 2160).

 

 

 The magnification chosen is important, if you go from a choose a higher magnification, the image size will jump to ten of thousands of pixels by tens of thousands of pixels.  Most image analysis software will not work with images this large. In practice, just zoom and center the image around the features you want and right click to export. 

 Use escape, Alt+F4, or the x button in the upper right corner of NDPView2 to exit the software.  This will bring you back to NDPScan.  Also, remember in Windows you can use Alt+Tab (hold Alt and keep pressing Tab) to cycle between full screen applications. 

 In NDPScan, choose to exit job (upper left) and then exit (lower left) to unlock the door and close the program. 

 

 

 

   

 Take your slides out and close the door.  Transfer your files to the core server, then log off.  You can leave the Nanozoomer on.  

   

  

Types of Labelling for Microscopy
The types of labelling used depends on the type of microscope or imaging technique. 

 1. Transmission Electron Microscope (TEM), Scanning Electron Microscope (SEM), Serial Block Face-SEM, etc. requires using heavy metals such as Osmium to label tissues. 

 2. Light Microscopes are used to image samples labelled based on Immunocytochemistry (ICC), Immunohistochemistry (IHC) and Immunofluorescence (IF), all of which utilize antibodies to provide visual details about protein abundance, distribution, and localization. 

 Labelling For Light Microscopes 

 What samples are used in ICC vs IHC? Sample Type and Preparation - tissue vs cells – As the name implies, IHC detection refers to tissue immunostaining, of either formalin fixed paraffin-embedded (FFPE) or frozen tissue. ICC refers to the staining of isolated or cultured intact cells where samples may be from tissue culture cell lines, either adherent or in suspension. 

 Immunocytochemistry (ICC) 

 Immunocytochemistry (ICC) is an immunostaining technique that detects antigens in cultured or primary cells via light microscopy. Fluorochromes are the most common reporter molecules in microscopy and are standardly used in multicolor ICC experiments. 

 . 

 The example above depicts a multicolor ICC/IF scheme for indirect protein detection. HeLa cells were probed with anti-Ki67 (nuclear protein) and anti-Alpha-tubulin (microtubular protein) followed by detection with DyLightTM488 (green) and DyLightTM550 (red) conjugated secondary antibodies, respectively. Fixed cells were counterstained with the nuclear dye, DAPI (blue) 

 Immunohistochemistry (IHC) 

 Immunohistochemistry (IHC) is a technique that uses antibodies conjugated to enzymes that catalyze reactions to form detectable compounds to visualize and localize specific antigens in a tissue sample. The root “histo-” specifically applies to biological tissue, so the process is only immunohistochemistry if it is being done in organic tissue. In contrast, immunocytochemistry applies the same process to individual cells. 

 First, the tissue and cells must be “fixed” using a chemical like formaldehyde. This stabilizes the structural properties of cells, preventing them from changing throughout the process. Next, cells need to be permeated using a detergent such as Triton X, which allows antibodies to enter the tissue and bind to epitopes within the cell. 

 Primary antibodies against a protein of interest are added, and secondary antibodies with enzymes like horseradish peroxidase (HRP) conjugated to their Fc domain are added to target the primary antibody. Enzymes like HRP can target certain substrate molecules like diaminobenzidine (DAB) and catalyze oxidation which results in the creation of a colorful compound. 

 This colorful compound will stay localized to the area the antibody was targeted to, staining the area near the protein of interest a different color from the rest of the tissue. Lastly, tissues are counter-stained using a dye like a hematoxylin to create contrast between the tissue stained using IHC and the non-colored regions for better visualization. 

 

 Immunohistochemistry (IHC) has found numerous applications in medicine, especially in cancer diagnosis. Lymphomas are among the cancers most dependent on IHC for correct diagnosis and treatment decisions. 

 Direct vs indirect immunofluorescence (IF) 

 ​​Immunofluorescence (IF) or cell imaging techniques rely on the use   antibodies  to label a specific target antigen with a fluorescent dye (also called fluorophores or fluorochromes) such as fluorescein isothiocyanate (FITC). Primary conjugated antibodies that are chemically labeled to fluorophores are commonly used in IF. 

 We distinguish between two IF methods depending on whether the fluorophore is conjugated to the primary or the secondary antibody: 

 Direct IF uses a single antibody directed against the target of interest. The primary antibody is directly conjugated to a fluorophore. 

 Indirect IF uses two antibodies. The primary antibody is unconjugated and a fluorophore-conjugated secondary antibody directed against the primary antibody is used for detection. 

 

 Labelling For Super-Resolution Microscopes 

 Super-resolution fluorescence microscopy, also called “nanoscopy,” enables the visualization of cellular structures beyond the diffraction limit of light. However, unlike electron microscopy, whose application is limited to fixed cells, nanoscopy enables live-cell imaging to study cellular dynamics in unprecedented spatial detail. 

 Green fluorescent protein (GFP) and its spectral variants have revolutionized biology, as they allow genetically encoded labeling, but they possess mediocre photophysical properties, generally emitting fewer photons than the best organic dyes by one or two orders of magnitude. 

 For super-resolution imaging, organic fluorophores are highly appealing because of their brightness and photostability . Organic fluorophores can be attached to proteins by combining click chemistry with unnatural amino acid incorporation. A second option is the direct coupling to proteins in live cells by using self-labeling proteins such as SNAP-tags (or a variant called CLIP-tag;  and HaloTags. Alternatively, labeling can be achieved by combining click chemistry and self-labeling proteins. Like GFP, these self-labeling SNAP-tags and HaloTags can be expressed as fusion proteins and selectively reacted with the substrates benzylguanine (BG) and chloralkane (CA), respectively, which are tagged with organic fluorophores. 

 Super-resolution techniques such as STED, PALM, dSTORM and Minflux localizes one molecule at a time. Single molecule localization techniques use ON/OFF principle. 

 

 STED uses a beam that turns fluorescence on, and another beam, shaped like a “doughnut” that turns fluorescence off allowing single molecules to be detected as it scans across the sample. 

 

 PALM/STORM uses same on/off concept but turns molecules individually off and on using photoactivatable proteins 

 Minflux combines STED and dSTOM or PALM principles in imaging. 

 

 

 Live Cell Imaging (similar to PALM/STORM) – no blinking buffer needed 

 Fixed or Surface Marker Live Cell Imaging (similar to dSTORM) – may require blinking buffer 

 Snap-tags, a self-labeling protein tag commercially available in various expression vectors that can be fused to any protein of interest and further specifically and covalently tagged with a suitable ligand, such as a fluorescent dye (snap-tag 647 is recommended). 

 New England Biolabs sell substrates that label the SNAP-tag® without the need for additional enzymes. Cell-permeable substrates (SNAP-Cell®) are suitable for both intracellular and cell-surface labeling, whereas non-cell-permeable substrates (SNAP-Surface®) are specific for fusion proteins expressed on the cell surface only. 

 Size of Labels 

 

 A single IgG is about 15nm and so the IgG complex could be >25nm 

 In contrast, VHH nanobodies are ~3nm 

 Proteintech sells ChromoTek SNAP/CLIP-tag VHH, recombinant binding protein 

 Minflux General Labelling 

 

 

   

   

   

   

  

X5000