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Phys598BP Spring 2016 University of Illinois at Urbana-Champaign

ANSWER KEY

Lab 2 (IGB): Bright Field and Fluorescence Optical Microscopy and Sectioning Location: IGB Core Microscopy Facility Microscope: Zeiss Axiovert 200M with Apotome Lab Instructor: Jaya Yodh, jyodh@illinois.edu Questions 1) What is the purpose of Köhler Illumination in light microscopy? (5 points)

Historically, this method was developed in 1893 by August Kohler to address the problem created by the use of filament bulbs as a light source. The latter often projected onto the sample creating regions of unequal brightness, thus interfering with critical illumination of the light source onto the specimen. Kohler illumination allows for independent control of the illuminating and imaging ray paths. The illuminating filament in the aperture set of conjugate planes does not disturb the image which lies in the field set of conjugate planes within the image forming path.

Modern microscopes are equipped to carry out Kohler illumination by using a collector lens to focus the lamp image onto the plane of the aperture (field) diaphragm of a condens er positioned at sufficient distance from the field lens. Controlling the opening of the condenser aperture thus controls the angle and amount of light rays reaching the specimen, and the light rays that pass through illuminating the specimen are brought to focus at the back focal plane of the objective. Here the images of condenser aperture diaphragm and the light source are brought into focus. Kohler illumination leads to optimum illumination of the specimen with uniformity in brightness and no glare. Furthermore, since the light source is not focused at the level of the specimen, the light at the specimen level is not grainy, is extended, and not compromised by dust and imperfections in the condenser lens.

2) What are the advantages of DIC relative to phase contrast? (5 points)

Phase contrast and differential interference contrast (DIC) micrsocopy are complementary methods that enable visualization of unstained transparent, non-absorbing (phase) specimens that are not observable by the human eye nor under brightfield illumination. The key to these techniques is the ability of phase specimens to modify the optical path difference between waves passing thru the specimen and those passing thru the surrounding medium.

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Frits Zemike won the Nobel Prize in1953 for building the first phase contrast microscopy in 1938. The basic principle was to create interference in light patterns that result in image contrast by manipulating the speed of the illumination light path hitting the specimen and by using annuli (i.e. rings) placed in the lens as well as the beneath the condenser in order to reduce the light wavelength by a ½ phase. The result was separation of the undiffracted from diffracted light such that contrast within the image became visible. Very dense regions have large optical path lengths and appear darker than background whereas regions that are less thick or with a refractive index less than the surrounding medium look much lighter relative to background.

George Nomarski developed differential interference contrast (DIC) microscopy in 1952 as an improvement over Phase contrast. The contrast in DIC is achieved by measuring the differences refractive index gradients across regions of varying specimen thickness. Illuminating light first passes thru a polarizer placed between the light source and the condenser. The resultant polarized light beam enters a 2-layered Wollaston prism which splits the light into two spatially separated beams. As the 2 beams pass thru the specimen, their wave paths are altered according to thickness, slope, and refractive index gradients of specimen details in proximity to each other.. After passing thru the objective, the beams are combined thru a 2nd Wollaston prism and then an analyzer which reconstitutes the vibrational planes of the beams which causes amplitude (path length) differences that are visualized as differences in brightness. This technique allows for greater depth of focus such that thicker specimens can be observed under higher magnifications. In a monochromatic image, areas of the object appear brighter as the optical path of the transmitted light increases and darker in areas of decreased length of optical path. In addition, steep gradients in path length generate excellent contrast, and images display a pseudo three-dimensional relief shading that is characteristic of the DIC technique

Advantages of DIC over phase contrast include the following: • DIC allows for imaging of thicker unstained specimens • DIC allows for use of higher numerical apertures and thus provides higher resolution • DIC does not produce the halo effect observed along edges in phase contrast • DIC imaging can be used in conjunction with fluorescence microscopy a better fluorescence image and

to pinpoint specific areas on a specimen before switching to the fluorescence mode to further examine the object.

• In DIC, the specimen appears bright on a dark background versus in phase contrast where it is dark on a light background. The former allows setting the condenser aperture at full-width so there is maximum illumination for focusing.

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3) Explain the meaning of the three curves on the following graph for the FITC filter set. (5 points) The graph below shows a typical emission output for a fluorescence filter cube or block that is housed in

a epifluorescence microscope setup. The filter block cube has the following components: 1) An excitation filter that transmits illuminated light only in the range of wavelengths that will excite the dye of interest. Currently, bandpass filters are used which have a well-defined short wavelength cut-on and long wavelength cut-off. 2) A dichroic beam splitter is a coated glass set at a 45 degree angle to the optical path of the microscope. The coating allows it to reflect the selected range of excitation light into the objective (and specimen) and then transmit the emitted fluorescence from the sample and allowing it to reach the detector. 3) An emission or barrier filter will attenuate all the light transmitted by the excitation filter while efficiently transmitting fluorescence (longer wavelength than excitation color) emitted by the specimen. Most microscopes have a turret that can hold from 2-4 filter blocks. The filters and dichroic beam splitters in the filter block are typically a matched set and should not be mixed

The graph shown in the handout is for Fluorescein isothiocyanate (FITC) which has an excitation and emission spectrum peak wavelengths of approximately 495 nm/519 nm (similar to Alexa 488). The blue peak (left) corresponds to the excitation wavelength range allowed thru the excitation filter (in this case, center wavelength is 480 nm with a full bandwith of 40nm). The red peak (right) corresponds to the emitted fluorescence wavelength range transmitted to the emission filter (in this case the center wavelength is 535 nm and full bandwith is 50nm) from the dichroic beam splitter. The green curve represents the dichroic mirror designed in this case with a cut-on wavelength of 505 nm where transmission is 50%.

The graph on the right shows another illustration of the excitation and emission spectrum overlayed on top of the excitation and emission filter cut-offs. Note the dichroic mirror eliminates the wavelength range where the excitation and emission spectrum overlap (i.e. stokes shift region)

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4) Include the following images and accompanying explanations: (1) Compare as a set (5 points)

• Bright field image of BAPE (Fluo) cells • Phase Contrast Image of BAPE (Fluo) cells • DIC image of BAPE (Fluo) cells

Note that you cannot see these thin transparent cells in bright field at all. With phase contrast in 10X objective, you see the nuclei features well however the image and cellular outlines are compromised by halo effect around the nuclei. In the DIC image under 20X objective, the pseudo-3D image of the nuclei (which is thicker region) is apparent although the fine details are not observable and the rest of the cell is flat.

(2) Fluorescent images (3 channels + merge) of BAPE (Fluo) Cells (compare as a set) (10 points)

• Discuss the differences between the wide-field fluorescence images of the FluoCells in 2D (what is labeled in each picture & characteristics of the fluorophore it is labeled with).

The FluoCells from Invitrogen are bovine pulmonary artery endothelial (BAPE) cells stained with with a combination of fluorescent dyes. Mitochondria were labeled with red-fluorescent MitoTracker® Red CMXRos. After fixation and permeabilization, F-actin was stained using green-fluorescent Alexa Fluor® 488 phalloidin; and nuclei were stained with blue-fluorescent DAPI. The excitation and emission wavelengths for the 3 dyes are listed in the table below. Thus, 3 different fluorescence filter blocks were used corresponding to the 3 dyes.

Phase Contrast (10X)

DIC (20X) Bright Field (10X)

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(3) Compare as a set and discuss differences between thin and thick specimens.(5 points)

o Color Bright-field Image of Plant Cross Section o Color Bright-field image of Pig Intestine

White balance and shading correction functions were applied to these images to make the background white and the shading even through the image, respectively. The pig intestine is stained with H&E Hematoxylin (H) and eosin (E) histological stain. H stains nuclei and E stains other regions of the cell. The pig intestine is only 6 µm thick and you can see the level of detail resolved for this sample in color brightfield. The plant stem is 40 µm thick and it is hard to focus on one plane and the resolution is much less compared to the pig intestine.

Plant intestine Plant stem (40µm thick)

Alexa488 DAPI

MitoRed Merge

Pig intestine (6 µm thick)

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(4) Wide-field fluorescence Z-stack image of plant cross section. Render 3D image and deconvolute using Autoquant software. (15 points total)

Note – in 2016 we did not do the Z-stack with the 20X objective but I include sample images at both magnifications in the answer key (plus more than 10 iterations just for you to see.

• Choose a mid-stack focused plane (slice) to display before and after deconvolution (10 iterations) for both 20X and 63X oil immersion objectives. Note the higher resolution at 63X oil compared to 20X.

• If time permits, also try increasing # iterations during deconvolution and display those for comparison. Doubling # iterations did not produce much difference in deconvoluted image with 10 iterations

Gallery of Z-stack From WF fluorescence

20X objective 96 slices (48 is boxed)

Deconvolved 10 iterations z-stack (mid slice 48)

Not deconvolved z-stack (mid slice 48)

Deconvolved 20 iterations z-stack (mid slice 48)

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Note – I did not ask for this in the lab report but thought I’d include examples for your benefit (some of the groups used the 5D viewer to get these 3D reconstructed images.

Gallery of Z-stack From WF fluorescence

63X oil 104 slices (53 is boxed)

10 iterations z-stack (mid slice 52)

20 iterations z-stack (mid slice 52)

Not deconvolved z-stack (mid slice 52)

3D reconstructed deconvoluted images

63X oil

20X

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• Explain in general terms what the deconvolution algorithm does to the original image Deconvolution refers to computational image processing tools that are used to improve contrast and

resolution in microscopy digital images by either removing or reversing blurring induced by the limited objective aperture. Typical digital images that are subject to deconvolution are 3D reconstructions made from a set of optical sections. In our lab we applied deconvolution to images taken in wide-field fluorescence which allows blurred light to reach the detector after which it is either subtracted or reassigned back to a source. In contrast, confocal microscopy places a pinhole aperture between the objective and detector which prevents out of focus light from reaching the sample.

In general, an image is formed by a convolution of the specimen with the point spread function, and for a large-matrix set of data (such as a 3D stack), if the image and point spread function are transformed into Fourier space, the convolution of the image by the point spread function can be computed simply by multiplying their Fourier transforms. The resulting Fourier image can then be back-transformed into real three-dimensional coordinates.

Deconvolution reverses the convolution process and attempts to reconstruct the specimen from a blurred image. The point spread function can be defined either theoretically by utilizing a mathematical model of diffraction, or empirically by acquiring a three-dimensional image of a fluorescent bead. Deblurring Algorithms are two-dimensional: they subtract away the average of the nearest neighbors in a 3D stack. For example, the nearest-neighbor algorithm operates on the plane z by blurring the neighboring planes (z + 1 and z - 1, using a digital blurring filter), and then subtracting the blurred planes from the z plane. While these algorithms are computationally simple, the can add noise, reduce signal, and sometimes distort image. Image Restoration Algorithms operate simultaneously on every pixel in a 3D image stack. Instead of subtracting blur, they attempt to reassign blurred light to the proper in-focus location by reversing the convolution operation inherent in the imaging system. The Adaptive Blind Deconvolution algorithm we used in the lab used the statistical techniques of Maximum Likelihood Estimation (MLE) and Constrained Iteration (CI). It does not require a measured or acquired PSF, but instead iteratively reconstructs both the underlying PSF and best image solution possible from the collected 3D dataset.