In the field of cell biology, it is essential to clearly see cellular structures and other cellular components. In experiments involving cell and cell structure examination, light microscopes, fluorescent microscopes, and electron microscopes are used. Electron microscopes are extremely powerful tools; however, the sample preparation and operation of the microscope require extensive training. Therefore, only the theory of electron microscopy will be covered in this course. Light microscopes, on the other hand, are relatively simple and are often used even at the introductory level of biology.

There are three elements needed to form an image: 1) source of illumination, 2) specimen to examine, and 3) system of lenses that focuses illumination on the specimen and forms an image. In a light microscope the source of illumination is visible light, the lenses are glass, and the image is viewed directly or focused on a detector (film or digital camera). An electron microscope uses a beam of electrons (the source is a tungsten filament) for illumination, electromagnets as lenses, and the image formed by the electron beam focusing on a fluorescent screen or a detector (film or digital camera).

Before beginning a discussion of microscopy, there are some important terms that should be reviewed; wavelength, resolution, and magnification.

Wavelength is the distance between the crests of two successive waves (figure 1). The ability of an object to disrupt a wave motion depends on the size of the object in relation to the wavelength of the motion. This means that the wavelength of the illumination source limits how small the object can be in order to be seen, so using visible light size must be greater than 300 nm, and 2 nm with an electron microscope. This is called the limit of resolution.

Resolution is defined as the ability to see two points that are close together as two separate points. A high resolution means that you can resolve much more detail, giving you a higher quality image, but a low resolution is limiting. The resolution itself is limited by the wavelength of incoming light and the numerical aperture quality of the lens system. Even with such limitations, a high quality microscope is still able to increase the resolution of an image 1000X that of the naked eye.

Magnification is an increase in the size of an image. Although an image can be increased up to 1000X the size of the actual object using higher magnification lenses on most modern microscopes (and about 100,000X with an electron microscope), the quality of that image is still limited by the resolution. Therefore, as you increase the magnification of your sample, the resolution of the image decreases, thereby decreasing the quality of that image.

In this laboratory, we will be using a binocular compound light microscope. Binocular refers to the fact that there are two ocular lenses through which we view our specimen. Compound refers to the fact that there are multiple lenses involved in the magnification of the image. Light refers to the way in which the image is focused.

Excercise: Proper Procedure for Use of a Light Microscope

1. Always use both hands to carry the microscope: one on the arm and one under the base.

2. Before turning on the microscope, be sure that:

1. The lowest power objective is in place.

· Important! Check the objective before focusing.

2. The lenses are clean (wipe gently with lens paper only).

3. The stage does not have a slide from a previous use.

4. The light source is turned all the way down.

5. The microscope is plugged in.

3. Place our slide on the stage and be sure that it is locked into place. Use the mechanical stage control to center the specimen under objective lens.

4. Look through the ocular lens and begin to focus DOWN with the coarse adjustment knob.

5. When the specimen is nearly focused, the fine adjustment knob can be used to sharpen the image.

6. Most modern microscopes are parfocal. They are also typically paracentral. As we increase the magnification on our microscope, be sure to pay attention when changing the objective – do NOT let the lens touch the slide! We may need to increase the light slightly and fine focus the image.

7. Calculate the Total Magnification at which we are viewing our sample by multiplying the power of the ocular lens (usually 10X) by the power of the objective lens.

8. To maximize the resolution of our sample, adjust the condenser lens to the position closest to the stage and open the iris diaphragm completely. We can adjust the contrast of our image by closing the iris diaphragm slowly until the image is optimal.

9. When we are done with the microscope, be sure to return it to the proper location, in the proper condition (light off and/or turned all the way down, lowest objective, no slides left on the stage, lenses clean, cord wrapped around the base and dust cover in place).

Note: Should we need to bring the microscope closer to us after setting it on the bench, do not drag it across the bench top. Pick it up to move it.

The fluorescent microscope is a modification of a light microscope. The modifications allow for visualization of fluorescently stained or tagged structures or molecules within the cell. To understand how this takes place, we must first understand the nature of fluorescence.

Fluorescence is a process that begins with absorption of light by a molecule and ends with its emission. When an atom absorbs a photon (or quantum) of light of the proper energy, one of its elections jumps from its ground state to an excited state. The electron then loses energy and drops back down to ground level, emitting another photon in the process. This emitted photon has less energy and a longer wavelength then absorbed photon (Figure 2). This results in two spectra uniquely associated with each fluorescent molecule, a characteristic absorption and emission spectra (Figure 3).

The fluorescent microscope is constructed in such a way as to allow visualization of only the fluorescent emissions of a particular wavelength. This is accomplished by using a filtration system to exclude emissions at all other wavelengths. There are two filters, 1) the exciter filter between the condenser and light source, and 2) the barrier filter located prior to the eyepiece objective. The illuminating light passes through the exciter filter before it reaches the specimen, passing only those wavelengths (absorbance) that excite (or absorbed by) the particular fluorescent dye. The barrier filter blocks out this light and passes only those wavelengths (emission) emitted when the dye fluoresces. Dyed objects show up in bright color on a dark background.

The condenser lens focuses light on the specimen causing fluorescent compounds in the specimen to emit light of longer wavelengths. Both excitation light from the illuminator and emitted light from the specimen pass through the objective lens. As light passes through the barrier filter, the excitation wavelengths are blocked or removed so that only the emitted wavelengths are viewed.

A light source should also be there. The objective acts both as a condenser lens for the excitation wavelength and the objective lens for emission. The emitted light from the excitation wavelength is isolated by the beam splitter.

There are a number of fluorescent dyes available that can stain different cellular structures or components. For example: DAPI is a dye that will stain the nucleus. Each dye has a different excitation and emission wavelength which determines which filter should be used and what color the object will appear to be. DAPI can be excited at a wavelength of 358nm, and will emit light at a wavelength of 461nm. If we've stained our cells with DAPI, we would use the filter corresponding to its excitation/emission wavelengths, and the nuclei of our cells would appear blue under a standard fluorescent microscope. Use of these dyes allows for visualization of where certain proteins, cellular structures, or other cellular components are in the cell at a given time.

Some examples using fluorescence are:

1. immunostaining: antibodies are linked to a fluorescent dye and when recognized by and bind to specific molecules (antigens) may then be visualized, indicating sites specific to those antigens;

2. indirect immunofluorescence: primary antibody attaches to the antigen, a secondary antibody is then attached to the primary and a fluorescent dye to the secondary, creating an additive effect with increased sites for labeling;

3. recombinant DNA techniques fuse a DNA encoding Green Fluorescent Protein, a naturally occurring fluorescent protein, to a gene coding for a specific protein of interest, introduced into cells where it is expressed and produces fluorescence; and

4. detection of ions using fluorescent probes sensitive to certain ions, i.e. calcium.

Electron microscopes are much more powerful than light microscopes and allow us to view (in detail) structures that are too small to be seen with a light microscope. The reason electron microscopes can reach higher magnifications is because they can produce images with much higher resolution. Electrons are used rather than light to focus the image, and since electrons have a shorter wavelength than light, they have a greater resolving power.

There are two types of electron microscopy (EM): Scanning EM and Transmission EM. Scanning EM uses a beam of electrons to scan the surface of an object. Typically, the surface of the object is covered with an electron dense material (such as gold), so that when the electron beam scans over the surface, secondary electrons are produced and collected by a detector. The detector is then able to produce an image based on the electrons it receives. Transmission EM is very different than scanning EM. This type of microscopy is used to view very thin sections of a specimen rather than the surface. The specimen is typically embedded in plastic and ultrathin sections are cut from the embedded tissue.

The sections are mounted on a metal grid and can then be viewed with the transmission electron microscope. To create the image, an electron beam is generated and then focused by an electromagnetic condenser lens. The electron beam passes through the specimen on the grid. Areas on the grid that are covered by part of the specimen will block electrons from passing through and will appear darker than other parts of the sample.

1. Obtain the Human Blood slide.

2. Identify 3 different cell types (one granulocyte, one monocyte, one erythrocyte) from each slide.

Refer to Figure 4 to identify the cell types.

Adapted from the following: Reference: Warren Strober, National Institute of Allergy and Infectious Diseases, Bethesda, Maryland. Current Protocols in Immunology. John E. Coligan, Ada M. Kruisbeek, David H. Margulies, Ethan M. Shevach, and Warren Strober (eds.). John Wiley & Sons, Inc. Online Posting Date: May, 2001. Print Publication Date: March, 1997

We will now use the concepts that we have learned to view different cell types and accurately describe them. For this exercise, we will use a tool called a hemacytometer (Figure 5). The hemacytometer is a slide that sits on the stage of the microscope. It contains chambers with grids (Figure 5, 6) where a liquid culture of cells can be added. The size of the grids is known, so a density of cells in the culture can be determined using the hemacytometer (Note: The hemacytometer should never be placed on top of another slide to measure those cells. It should only be used with liquid cultures!). Before using the hemacytometer, clean the surface with lens paper, as well as the coverslip. The hemacytometer coverslip is specially made and is heavier than a conventional coverslip in order to overcome the surface tension of liquid. After the coverslip is placed on the counting surface, a cell suspension is added via the v-shaped well using a micropipette tip, filling by capillary action. Only add enough liquid to cover the counting surface. Place the hemacytometer on the microscope stage and bring the counting grid into focus at low power.

The hemacytometer grid is divided into 9 large squares, each with a surface area of 1 mm² and a depth of 0.1 mm. Therefore, if we count the total number of cells in the four large corner squares plus the middle square, we will have the cell number in a 0.5 mm² area. There are 1000 cubic millimeters in one cubic centimeter (which is equal to a milliliter). We may then calculate the number of cells using the hemacytometer, use the following formula:

(average number of cells/mm² square) x (10) x (1000) = number of cells per milliliter

The hemacytometer grid is divided into 9 large squares (one large square is shown in red), each with a surface area of 1 mm² and a depth of 0.1 mm (when coverslip is in place). Therefore, if we count the total number of cells in the four large corner squares plus the middle square, we will have the cell number in a 0.5 mm³ area. There is 1 mm³ in 0.001 cm³ (which is also equal to 1 µl). We want to determine the cell concentration in 1ml of suspension, so we use the metric conversion: 1ml=1000µl.

We can then calculate the number of cells per ml of solution using the hemacytometer, by incorporating the following formula:

(average number of cells/mm² square) x DF x (10) x (1000) = number of cells per milliliter

From the equation above:

DF: Dilution factor

10: to correct for the depth under the coverslip

1000: to correct for the conversion from microliters to milliliters

Example: Suppose that we counted a total of 125 cells in five large squares. We have an average of 25 cells/mm2square. Multiply by 10 to correct for the area from hemacytometer surface to coverslip. Again, multiply by 1000 to determine cell count per ml (250,000).

If our cell number is large, dilute the cells, taking care to mix the cell suspension prior to taking an aliquot. We will then need to correct for that dilution by multiplying our cell number by the dilution factor. Also, if we are using a staining solution, we need to correct for that dilution.

Example: Using the above example, suppose we had diluted our cells by a factor of 10, or a 1 to 10 dilution, we would then multiply the final cell number by 10 and our answer would by 2,500,000 cells per ml in the original cell suspension.

Article “The HL-60 promyelocytic leukemia cell line: proliferation, differentiation, and cellular oncogene expression” by S. J. Collins (1987) Blood 70, 1233-1244.

We will be using the cell line HL-60 as our model for this course. The HL-60 cell line was established as a long-term suspension culture of human myeloid leukemic cells (Collins et al, Nature 270:347, 1977). The cell line was originally derived from a patient with acute promyelocytic leukemia and subsequently successfully maintained continuously in tissue culture as a suspension culture. This success was remarkable not only because this cell type is difficult to culture, but also because the cells could be grown continuously in suspension cultures. Factors critical to the ability of HL-60 cells to proliferate in vitro include: 1) insulin receptor expression; 2) transferrin receptor expression; 3) colony-stimulating factor stimulation. Because the HL-60 cells can be grown in culture for a continuous period they are considered an excellent stem cell-like model. True stem cells are difficult to maintain in culture making experimentation with them inconsistent. The capability of inducing differentiation in HL-60 cells is also of experimental significance, allowing for studies of biochemical pathways in the hematologic lines to be explored.

Important HL-60 characteristics

Promyelocytic leukemia cells continuously proliferating in suspension culture (our doubling time is about 22-24 hours) – unusual human myeloid leukemia.

• Factors contributing to adaptation to continuous growth (stem cell-like) are

  • 10 cell surface expression of transferrin receptors;

  • cell surface expression of insulin receptors;

  • production of colony-stimulation factors.

Both transferrin and insulin are required to be available in medium supplementation. Transferrin is required for an active iron transport system for cell proliferation to take place. As cells are induced to differentiate, transferrin receptor expression decreases. Insulin receptor expression decreases when HL-60 cells are induced to differentiate into granulocytes. Normal myeloid progenitor cells are stimulated to grow in culture by the colony-stimulating factor (CSF) compounds. HL-60 cells appear to produce CSF-like compounds that simulate this activity. There appear to also be some genetic abnormalities present in HL-60 cells that may play a role in allowing these cells to be grown continuously in vitro.

Ability to induce differentiation into four types of cells: granulocytes, monocytes, macrophage-like cells, and eosinophils. Below we list the cell type that cells can differentiate into followed by common compounds that elicit the differentiation:

• Granulocyes: Dimethylsulfoxicide, Retinoic acid, Actinomycin, Tunicamycin

• Monocytes: Vitamin D3, Sodium butyrate, TNF (tumor necrosis factor), Ara C

• Macrophage-like (very similar to monocyte): PMA, TPA (12-O-tetradecanoylphorbol-13-acetate), Teleocidin

• Eosinophils: Butyric acid, GM-CSF, alkaline media(pH 7.6 to 7.8)

For next class, you’ll need to read the Xie et al paper.

Procedure:

1. Read the article “Fibronectin-mediated Cell Adhesion Is Required for Induction of 92-kDa Type IV Collagenase/Gelatinase (MMP-9) Gene Expression during Macrophage Differentiation” by Bei Xie et al. (1998) J. Biol. Chem. 273 (19), 11576-11582.

2. Can you identify the hypothesis, experimental design and conclusions? Briefly describe these sections.

3. Are the conclusions of this article supported by the data? Do they answer the original question?

4. Locate and name the author(s), the date the article was published, and the journal the article was published in.

5. Now, try to find a review article that is related to this topic. What kinds of differences can you see between the two types of article?

6. Define any terms you are not familiar with. In particular, define the following terms: tumor necrosis factor, cytokine, colony-stimulating factor, matrix metalloproteinases, extracellular matrix, fibronectin, integrin

Reading a Scientific Paper: Discussion of Xie et al. (1998) J. Biol. Chem. 273 (19): 11576-82.

Title

• Should be informative and should provide the main focus of the research presented.

• This will be the first thing someone looks at to determine whether or not they want to read the paper – often very detailed for this reason )you want for your paper to be cited as often as possible!!)

• What does the title of this paper tell you? What are some of the key words?

Abstract

• This should be a one paragraph summary of the paper.

• It should include the major findings and the significance of those findings.

• This is the second thing a person will read to help them decide if they want to read the entire paper – it should be informative and provide the “highlights” of the research.

• Does this abstract tell you the purpose? What is it?

• What are the major research findings presented in the abstract?

• What is the significance of the work as presented in the abstract?

Introduction

• This section provides the background necessary for someone to understand the paper.

• Generally the authors assume that the audience has a base level of knowledge, so many terms will not be explained. Note: When reading these papers, if you come across a term you don’t know, you should always stop and look it up to aid in your understanding of the paper.

• The following terms may not be familiar to you, but this paper assumes you know what they are. Look them up and provide a brief explanation (1-2 sentences) for each: Tumor Necrosis Factor, Cytokine, Colony Stimulating Factor, Matrix Metalloproteinase, and Integrin.

Materials and Methods

• There are different ways to present the Materials and Methods – sometimes this section comes right after the introduction (as you would do for a lab report), but sometimes it is given at the end of the paper, just before the references.

• Most people in the scientific community will not usually read the M&M as they will the rest of the text. If there is a question about how an experiment is performed or if a research wants to do similar experiments, they will consult this section, but it is not usually read otherwise.

• The M&M should provide enough information to allow you to go back and repeat the experiments and get the same results.

• Often, a long experimental procedure that has been previously published will be cited instead of being written out – Example: “the percentage of cell adhesion ad spreading was determined as described previously (24)” – from page 11584.

Results

• In any paper you read, you should be able to get all the information you need from the text alone OR from the figures + legends alone, for example: Consult Figure 1 – compare to first paragraph of the results section.

• The results section is simply a statement of the results – there should be very little (if any) discussion of what the results mean in the context of other work in the field. This means you will not find many citations/references in the results section.

• Remember, the objective of the results section is just to convince the reader that the result is REAL.

Discussion

• The discussion generally presents a brief summary of the problem/hypothesis, followed by a summary of the results.

• The results are generally discussed in the context of current literature (the work of other scientists in the field).

• Finally, the last paragraph should summarize everything and include the significance or importance of the research that was just presented. This paragraph will often look a LOT like the abstract.

• Does the discussion help to clarify the significance of this paper for you? How?

References

• This is where all of the references are listed. This section is generally pretty lengthy, including all of the sources that were cited in the intro, methods, and discussion (maybe a few from the results section too).

1. Barrier filter: filter near eyepiece that blocks light entering system through exciter filter and only permits light being emitted by sample to be viewed

2. Binocular: two ocular lenses

3. Compound: objective and ocular lenses

4. Dye exclusion test: preparation of cells in a trypan blue solution to determine cell viability; viable cells with intact membranes exclude the blue dye

5. Exciter filter: filter near illuminator that permits lights through to the sample in order to excite its photons

6. Fluorescence: light emitted during the absorption of a certain smaller wavelength of higher energy

7. Hemacytometer: specialized slide used to determine the density of cells, which allows for a simple conversion of cells/mL

8. Limit of resolution/Resolution: the least separation of two images so that they are seen as separate when viewed through a microscope

9. Magnification: the ratio of the size of an image to the size of the object

10. Paracentral: object remains in the center of the line of view when the magnification is changed

11. Parfocal: lenses stay in focus when the magnification is changed, however a very fine adjustment may be made

12. Total magnification: the power of the objective lens (4x, 10x, 40x, etc) multiplied by the power of the eyepiece, usually 10x.

13. Transmission EM: a high-energy beam of electrons is transmitted through a very thin sample to generate image