الميكروسكوب الضوئي مبعثر استنادا فلاتر غابور ثنائي الأبعاد

In this video, optical scatter microscopy is used to quantify mitochondrial fragmentation during apoptosis. Although optical scatter microscopy does not normally require fluorescence in this protocol, fluorescent images will be acquired for validation purposes. Following setup of the imaging system, cells are labeled with MIT tracker green and treated with stor sporin to induce apoptosis.

Optical data are acquired to detect changes in the orientation of cellular organelles within two hours. The onset of mitochondrial fragmentation can be measured using the optical scattered data, which shows a decrease in the degree of orientation of subcellular particles. The data are then validated by correlating the optical scattered data with the fluorescent regions containing mitochondria.

Hi, I'm Nada Bustani from the Bio Optics Laboratory in the Department of Biomedical Engineering at Rutgers University. I'm Rob Pak from the Bio Optics Lab, And I'm Deen also from Bio Lab. Today we'll show you a procedure for quantifying subcellular morphological dynamics using a method that does not rely on fluorescent dyes.

We use this technique in our laboratory to study dynamic changes in subcellular structure during apoptosis or program cell death. So let's get started. Begin this procedure by sterilizing gloved hands with 70%ethanol.

Then place a prepared 100 nano molar stock solution of miot tracker green and prewarm, BAEC cell culture medium in the hood with the light off and sterilize all surfaces. Next place a six well culture dish with cells plated on cover glass under the hood. Using a paste pipette connected to a vacuum line.

Remove the cell medium from the wells. Then immediately add two milliliters of the labeled medium to each well to label cells.Mitochondria. Cover the tissue culture plate to prevent exposure to direct room light and quickly place the cells back in the incubator.

Incubate for 45 minutes at 37 degrees Celsius while the cells incubate. Prepare the optical setup first turn on the mercury arc clamp. Then turn on the computer microscope cameras and laser plug in the digital micro mirror device or DMD in the spinning diffuser.

Next, look through the microscope eyepiece to see whether the optical launch is aligned. The field of view should be brightly illuminated by laser light. To load the sample, place one to two small drops of immersion oil over a 63 x oil immersion objective while it's all the way down.

Then on the stage mount a slide consisting of a grid of lines at regular 10 micrometer intervals called the gradi, and raise the objective so that the oil grabs it focus. Next, focus the hexagonal edge of the condenser field. Stop so that it's aligned in central Kohler illumination.

The hexagonal edge should be centered in the eyepiece and sharply focused. Then using the condenser field stop position knobs center, the condenser field, stop over the field of view. Start IP lab and input the settings to operate the camera.

Confirm that the camera is set to frame transfer mode. Run the acquire focus command to start live preview. Then set the index prefix and the file location to which the images will be saved.

Next, initiate RS image. Designate the settings to operate the cool snap camera. The clocking mode should be set to normal.

Start the DMD software. Place the dark field iris on the script menu. Then click on load and reset.

To run the script manually set the microscope optiv bar in viewport to LSM. This will send the image through the DMD in the aligned optics. Projecting the dark field image onto the CCD of the Cascade five 12 camera.

The dark field image should appear on the live preview in IP lab. If necessary, focus the image on the live preview. Set the exposure time high enough to ensure at least 10, 000 counts of signal in the image.

Then take a snapshot of the field of view using the acquire single command after acquisition. Use the save as indexed command to save the image to disc. The image of the GTI measures the size of the field of view.

Now move the GTI sample so that it's beyond the field of view and only the background is visible. Capture a background image of the field with an exposure time sufficient to acquire at least 5, 000 counts. This image will aid in background subtraction of the unfiltered images.

Now to acquire Gabor filtered images of the background load the Gabor Filter Bank script to the DMD control software. Run the entire script to buffer the filters to the onboard memory of the DMD. This may take a few minutes once the entire script is buffered.

Use the start and stop markers within the DMD software to ensure that only one set of filters corresponded to one Gabor Like filter is loaded at a time. Run the script to acquire filtered images of the background. The live image preview should change from dark field to the filtered image.

For that filter, open the acquisition script in IP lab from disc. Adjust the exposure time to ensure that at least 2000 counts are acquired. As the DMD script is running, cancel the live preview in IP lab and run the acquisition script.

This will automatically acquire index and save the filtered image to disc. Once the first image is acquired, stop the script running in DMD. Delete the used commands from the script.

Then replace the start and stop markers at the beginning and end of the next filter set. Repeat the acquisition in IP lab until the entire filter bank has been used and all the filtered images have been acquired and saved. Prepare to plate the cells by plugging in a soldering iron on the lab benchtop and heating the L 15 medium to 37 degrees Celsius.

Make a workstation with a paper towel and a Kim wipe to absorb any spilled medium and to keep the area clean. Prepare several wicks by tearing and twirling kimm wipes. The wicks will be used for drawing the fluid through the cell plate.

Next, use the metal sample holder to create a cell sandwich. Use a syringe to apply a thin bead of vacuum grease around the upper periphery of the hole in the machined metal plate, extending about halfway to the ends of the grooves on each side. Gently pass a clean number one cover slip onto the grease.

Flip the plate over and apply the grease around the hole. Turn off the room lights wearing ethanol sterilized NI trial exam gloves. Retrieve the cells from the incubator.

Remove the cover slip that will be used for the experiment from the six Well plate. Carefully dry the cover slip. Then press the cover slip cell side down into the greased metal plate over the viewing hole.

Sandwiching the cells within the plate. Ensure that no air gaps remain, and that the grease has formed a watertight seal so that medium can be loaded. Now flip the plate back over pipetting 200 microliters at a time.

Add prewarm L 15 medium into the sandwich through the groove between the upper cover slip and the metal plate until the liquid almost reaches the groove on the other side. Next, wash the cells by holding a wick at the opposite groove and pipetting an additional 200 microliters of medium into the sandwich. Medium will flow from one side to the other as the wick removes old medium.

Be careful to prevent any bubbles from forming within the liquid during this step. Repeat this process two to three times using a new wick for each rinse. Now invert the plate once more supporting the plate from the edges so as to trap the liquid in the cell reservoir.

Dip the soldering iron into the veap beaker to quickly melt some of the veap, which will then clinging to the soldering iron tip. Using the soldering iron tip carefully apply the molting veap around the edges of the bottom cover slip. Continue dipping and applying all the way around the cover slip perimeter, sealing it to the metal plate.

This creates a good mechanical seal so the cover slip won't detach from the plate as it's being viewed. Clean any residue from the bottom. Cover slip surface by bawling up a kim wipe and cleaning the cover.

Slip in a single sliding motion to ensure that the cover slip is cleanest in the center where it'll be viewed. Return the six wall plate to the incubator. Take the plated cells to the optics lab and mount on the objective.

As before, find a field of healthy looking cells and acquire a dark field image of the field of view. Then align the microscope in DIC and making sure the exposure times are adequate. Acquire a DIC image with a cascade camera.

In this setup, fluorescent images are acquired using the cool SNAP camera while the microscope is still aligned in DIC. Set the microscope VAR to one x and viewport to 100%binocular and divert the image from the eyepiece to the camera. Place a blue LED over the condenser to illuminate the field.

Then preview the field of view in RSS image and adjust fine focus. If necessary, acquire A DIC image and save to disc. Note how the field of view is different from the one obtained from the cascade camera.

These images will have to be registered during the analysis phase of the experiment. Next, move the fluorescent filter cube into position and acquire a fluorescence image by briefly turning on the fluorescence excitation using the microscope shutter, and then turn it off as soon as the acquisition is completed. To minimize photobleaching, save the fluorescence image to disc.

To acquire filtered images, reset the microscope to field and send the light through the LSM port. As before, run the entire Gabor filter bank script as before to complete the data acquisition for one time point while the cells are still on the stage and without disturbing the field of view. Use the wicking method to switch out the regular L 15 medium for the same containing a one micromolar solution of STS.

During the course of the experiment, add additional medium to prevent drying out by pipetting medium into the groove of the cell plate without removing it from the stage or disturbing the field of view. Now, repeat these steps for subsequent time points until the experiment is completed. The collected data will include a large number of filtered images that must be processed to extract the subcellular structural data.

Here two example experiments are shown. This figure shows sequential images of a marine diatom algae sample. The oriented features are clearly visible.

In dark field imaging. The optically filtered images labeled five equals zero degrees to five equal 160 degrees are shown alongside the unfiltered image of the sample. For comparison, the set of nine Gabor filtered images of the diatom were processed pixel by pixel for object orientation.

At each pixel, the maximum signal was divided by the average signal as a function of filter orientation to give an aspect ratio value. Here, an aspect ratio near one indicated by blue is present in areas in which there is no preferred response angle. Greater values indicated by red indicate areas in which a higher preferred angle response is present.

The brightness and code significance of the total GABOR filter response, the filter orientation, giving the maximum signal at each pixel corresponds to the direction perpendicular to the underlying object orientation. Substructure particle orientation is encoded in a quiver plot where each line closely agreed with the underlying local object orientation visible in unfiltered dark field. The quiver plot shows the orientation of objects with response intensities greater than 10%of maximum.

These dark field and optically filtered images of a field contained live bovine endothelial cells, filtered images were subsequently acquired every 20 minutes for a period of three hours after treatment with the apoptosis inducer stor sporin. Here, the aspect ratio map in the top panel and the fluorescence imaging of mitochondria in the bottom panel are shown as a function of time. The color hue represents the degree of orientation, as before these time plots compare the decrease in the degree of orientation of the particles aspect ratio in endothelial cells treated with s thoro sporin, the individual traces represent time plots within single cells.

The drop-in orientedness is confined to the regions of the cells that register with fluorescent regions. Now, the validation against fluorescence has been achieved. It becomes archival.

Thus, in the next experiments, the mitochondrial measurements can be made by omitting the fluorescence imaging steps and directly using the optical scatter method. We've just shown you how to monitor mitochondrial fragmentation during apoptosis using light scattering signatures based on Gabor filters. When doing this procedure, it's important to have a good lifestyle preparation and to ensure that the optical setup is well aligned and calibrated.

So let's take sense for watching and CoLab with UI experiments.