Background on immunofluorescence

The workshop

Whilst the nucleus and some other organelles are easily visualized using brightfield microscopy, most other sub-cellular structures cannot be differentiated or even seen using conventional microscopes. Therefore these structures and biomolecules must be demonstrated using more specific techniques such as immunofluorescence and fluorescence microscopy.

In this project you will be provided with a coverslip containing cancer cells grown in culture. You will then expose the cells to antibodies directed against proteins found in subcellular structures, allowing these primary antibodies to bind to those structures. You will then treat the cells with secondary antibodies directed against the primary antibodies. These secondary antibodies have been conjugated to a fluorescent dye, effectively attaching a fluorescent label to the structure you are interested in seeing. You will then use a fluorescence microscope to image the cells, taking images of each colour channel and then combining the images to create a merged image which demonstrates all structures.

Getting started

Before you begin, make sure that you are familiar with the relevant theory behind the techniques we will be performing. This manual contains several appendices which will provide you with this information. Make sure you read this information before proceeding.

Appendix A : DNA

Appendix B : Proteins

Appendix C : Using a micropipette

Appendix D : Glossary of terms

Information on other cell and molecular biology concepts and techniques are provided at the SPARQ-ed website here.

Theoretical basis of the project

Tissue culture and cancer cell lines

When researchers want to obtain sufficient quantities of organisms to use in their studies, they may grow them up in a vat of broth containing all of the nutrients they need to survive. This is called culturing the organisms, and for many simple, free-living organisms such as bacteria or fungi, the process is relatively straightforward. These organisms often require very little in the way of specialised nutrients and so long as the culture conditions are suitable, they will undergo almost constant division, increasing their populations at an exponential rate.

If cells from a multi-cellular organism like a human are to be used in research, the culture methods are not nearly so simple. Firstly, cells from multi-cellular organisms have differentiated to such an extent that they can no longer survive without the complex systems of nutrients and stimuli provided by the other cells in the body. As a result, growing human cells in culture requires the use of growth media containing a complex mixture of basic nutrients and specific growth factors provided by the inclusion of serum (the liquid component of clotted blood). In addition, body cells spend most of their time carrying out the functions which allow them to play their roles within the body. This means that they are not constantly dividing - in many cases they must be stimulated to enter mitosis, and in some cases do not divide at all. As a result, human cells in culture often proliferate very slowly.

One solution to this problem is the use of cancer cells. One of the hallmarks of cancer is the loss of regulation of the cell cycle, leading to cells constantly passing through division cycles. This results in hyperproliferation of the cells, which in the body leads to the growth of tissue masses known as tumours. Cancer cell hyperproliferation means that they can be grown in culture for many generations, with further cultures able to be sub-cultured from the original – they are effectively immortal. This has led to the production of cancer cell lines, each of which was derived from a tumour recovered from an individual with cancer.

Most types of cancer have numerous cell lines which researchers can use to study the cancer in question. Each of these cell lines have particular characteristics dependent on the cancer from which they were derived, allowing researchers to select the line which most closely matches the situation they are working on. Unfortunately, being cancer cells, the cells contain errors and abnormalities which mean that they do not often behave as normal cells do, making them inappropriate models for normal cells, or even for cancers of other parts of the body. In addition, the same errors which remove the cell cycle controls in cancer cells may also remove mechanisms which regulate damage to the DNA, allowing mutations to accumulate in the cells which make them even more distinct from the cells from which they were derived

HeLa cells are possibly the best known of all cancer cell lines. They were originally recovered from a cervical cancer removed from a patient named Henrietta Lacks in 1951. The cancer resulted from an infection by a strain of human papillomavirus (HPV18), and so the genome of these cells also contains the genome for this strain of HPV. HeLa was the first cell line to be successfully and continuously cultured in vitro and have been used widely around the world since the physician who first subcultured them made them and the techniques used to grow them freely available to scientists around the world. HeLa cells have played an important role in many important medical discoveries, including the development of the Polio vaccine.

In this workshop, you will be using a genetically modified strain of HeLa cells. These cells have a modified gene for one of the histone proteins in the nucleus (H2), where the gene includes a gene derived from a jellyfish (the gene for green fluorescent protein, or GFP). The histones are a family of proteins around which DNA wraps and which assist in the packaging of DNA and help to regulate the expression of genes. When the modified gene is expressed, the protein produced includes a GFP “tag” on the end which glows green when exposed to blue light. This has produced a cell line in which the nucleus fluoresces green, which means that this green signal can be used in place of a DNA dye such as DAPI. This fluorescence is produced in living cells, allowing them to be used to study the function and appearance of the nucleus at different stages of the cell cycle through live cell imaging.

Preparation of cells for immunofluorescence

Using routine brightfield microscopy, a reasonable amount of detail in the cell can be determined. Large structures such as the nucleus (and its nucleoli) and vacuoles are easily distinguished inside the cell with even fairly rudimentary microscopes. Some other structures can also be demonstrated with the careful use of dyes and stains which give a colour based on chemical reactions with cellular components, and this forms the basis of cytochemistry (on individual cells) and histochemistry (on thin sections of tissue). However, even these methods are not nearly specific enough to properly demonstrate the wide variety of subcellular structures and materials. Immunofluorescence is a technique which uses the highly specific binding of antibodies to their target antigens as a way of demonstrating materials and structures inside cells.

Before any immunofluorescence procedures can be carried out on the cells, they must be properly prepared. For this workshop, the cells have been grown on a coverslip placed on the bottom of a culture plate. When a new culture is needed, a suspension of cells is prepared with the cells floating around inside the cell culture medium. The number of cells per unit volume (generally per millilitre of medium) can be calculated using cell counters and a specific number of cells added to a dish containing culture medium by adding the correct volume of cell suspension (eg. if the density of cells in the suspension is 100,000 cells / mL and you needed 100,000 cells to start a culture, you would add 1mL of suspension). The exact seeding density depends on the reproductive rate of the cells you are using and how dense you want the culture to be in the time you have.

When the cells are added to the culture dish, they sink to the bottom and attach through proteins embedded in the cell membrane. Cells that have attached to the bottom of the dish take on the appearance of a fried egg, with the nucleus as the yolk and the cytoplasm as the white. Cells only release from the bottom of the flask when they are in the process of dividing, becoming spherical as they carry out mitosis. For this workshop, the cells have been added to a 6-well culture plate. In each well, a number of coverslips were added to the wells prior to the addition of the cells. When the cells settled to the bottom of the well, some landed on the coverslip. This means that the cells can be easily removed from the well and treated while on the coverslip, and eventually the coverslip containing the cells can be mounted onto a microscope slide for observation. The coverslips have been previously treated with Poly-L-Lysine, a peptide which acts as a cellular “glue” and assists in holding the cells (including mitotic cells which have rounded up) onto the coverslip.

At several times during the course of the immunofluorescence procedures, cells will need to be washed. This is usually done by replacing the liquid in the well they are contained in with phosphate buffered isotonic saline, a solution of sodium chloride at 9g/L in a buffer which keeps the pH at a physiological level of around 7.2. Because the cells are stuck to the coverslip, the liquid can be gently removed and replaced with a solution which does not put the cells under osmotic stress.

Once cells die (which starts to occur once their nutrients and ideal growing conditions have been removed), they start to break down and lose their structure and integrity. A typical live cell taken through the processes needed for immunofluorescence would have started the breakdown process by the time the techniques have been completed. Therefore the cells must be preserved prior to undergoing antibody treatment. This process is called fixation, and is the same technique used to preserve biological specimens in museums. Fixation relies on “freezing” the proteins in place using a chemical process rather than temperature. The tertiary structure of proteins is maintained by weak hydrogen bonds between the side chains of the amino acids that make up the proteins. These hydrogen bonds are easily broken by increases in temperature or changes in pH, resulting in a loss of protein structure called denaturation. Formalin is a buffered solution of the gas formaldehyde which forms permanent covalent crosslinks between protein chains and locks them into their tertiary structure, regardless of minor changes in temperature or pH, and thus preserves the structure of the cell. In this investigation, you will use a type of formalin called paraformaldehyde to fix the cells, although other fixatives like ice cold methanol can also be used.

Once cells have been washed, fixed in paraformaldehyde and washed again, they must then be permeablised. The antibodies used for immunofluorescence are large protein molecules which cannot cross the cell membrane, so the cell membrane must be removed. It is very important that the cells must be fixed prior to permeablisation, otherwise the cell will burst.

To understand why it is important to fix the cells, it may help to imagine the cell as a dome tent filled with cooked spaghetti (with the nucleus as a beachball inside it, perhaps). The shape of the cell is maintained by structural proteins such as Tubulin, represented in our analogy by the cooked spaghetti, while the whole shape of the cell is maintained by the cell membrane, represented by the fabric of the tent itself. If you were to cut away the tent fabric (removing the cell membrane through permeablisation), all of the spaghetti would fall out and spill onto the ground. Washing the cell would then have the effect of hosing the mess away.

Fixing the cells would be like drying the spaghetti out. It would shrink slightly and revert back to its state prior to cooking, however it would be in the same position it was in when the drying occurred. The tent fabric (cell membrane) could then be removed and you would be left with a mound of dried spaghetti in the same shape as the tent, but without the tent around it (see figure below). Chemical fixation does make some changes to the proteins in the cell, however enough of the original structure is often left to retain some of the proteins’ functions, including the ability of the proteins to bind antibodies and the fluorescence characteristics of green fluorescent protein (both of which are vital for this investigation.
 

In this workshop, cells will be permeablised using a detergent (Triton X100). Detergents work by solublising lipids in water, and since the cell membrane is made up of lipid molecules, the Triton X100 in this method removes the lipids from the membrane, allowing large molecules such as antibodies to enter the cell and bind to proteins inside. At the same time as they are permeablised, the cells will be blocked. Blocking involves the addition of a solution of proteins (usually bovine serum albumin, or BSA). Cells may contain proteins which can bind onto antibodies and other proteins non-specifically, rather than the antibodies specifically binding onto their target proteins. The BSA “soaks up” these non-specific binding sites, ensuring that the only way the antibodies can bind is via their specific targets. Once permeablised, blocked and washed once more, the cells are ready to be treated with antibodies.

Antibodies and immunofluorescence

Antibodies are large, complex proteins made by specific immune cells in the body (the plasma cells, which are derived from populations of B lymphocytes). Antibodies are produced in response to the presence of an agent in the body which the immune system recognizes as being not from the body. These agents are usually foreign materials, such as proteins on the surface of microbes, although in some cases the immune system may target the body’s own proteins, causing autoimmune diseases such as Type I Diabetes and Rheumatoid Arthritis. 

Antibodies are composed of four peptide chains (two light chains and two heavy chains) arranged like a capital letter “Y” (see figure at right). Most of an antibody’s structure is constant between antibodies (with some variation between different classes), however at the ends of the “arms” of the Y is a small, highly variable region made up of a region of the light and heavy chains. It is changes in the shape of this region which determines the difference between antibodies. The shape of this variable region is such that it fits around and binds very specifically to a region on the material it is raised against. The area which it binds to is called the epitope, while the material where the epitope is found is called the antigen. A different antibody is made in response to each epitope on each antigen, with the variable regions conforming exactly to each epitope. This means that antibodies are made which bind very specifically to particular epitopes on antigens. This specificity is so high that two antibodies can be generated which can tell the difference between two versions of the same protein, one which has a phosphate group attached and one which does not. This high level of specificity makes antibodies extremely useful in detecting particular proteins, and is used in techniques such as diagnosis of disease agents in the blood, western blotting, immunohistochemistry, and, of course, immunofluorescence.

Antibodies are made in animals, or in cell lines derived from those animals. To generate an antibody against a particular protein, samples of that protein are injected into a test animal. A better antibody response is generated if the target protein comes from a different species to the animal used to raise the antibody as the host animal would recognize the protein as being foreign. For example, if you are interested in using antibodies to detect a human protein, you would use a non-human animal such as a mouse, rabbit or goat to raise the antibody. Once the animal starts making the antibody, it can be recovered from the animal’s blood and purified for use. In some cases the lines of plasma cells which are making the antibody can be isolated and fused with a cancer cell, making a hybridoma cell line which has the immortality of the cancer cells and the antibody-generating properties of the plasma cells. This means that the antibodies can be produced in culture without the need for using further animals.

The technique you will be using is indirect immunofluorescence. Direct immunofluorescence involves chemically combining a fluorescent dye directly to the primary antibody raised against the protein you are interested in. The antibody, with its fluorescent label binds onto the target protein in the cell, essentially colouring the target protein with the label (see figure below). Direct immunofluorescence is not often used now, as it would involve making a range of differently labeled antibodies for every protein studied in laboratories. In addition, the sensitivity of the technique is low, as there is only one label for each binding site.

With indirect immunofluorescence, the primary antibody is left unlabelled. It still binds to the target protein, however it must be itself demonstrated using a labeled secondary antibody. This secondary antibody is generated in the same way as the primary antibody, however its target is antibodies from the species in which the primary antibody has been raised. For example, if the primary antibody was raised in a mouse, the labeled secondary antibody would be raised in another species against mouse antibodies (eg. goat anti-mouse). This allows laboratories to have a wide selection of primary antibodies directed against any protein they might be working on, and then a smaller selection of labeled secondary antibodies directed against the species the primary antibodies were raised in (eg. anti-mouse, anti-rat, anti-goat, anti-chicken, etc). In addition, because multiple labeled secondary antibodies can bind to a single primary antibody, the signal strength is higher, resulting in greater sensitivity (see figure below).

 

If primary antibodies from different species are used, then multiple proteins can be targeted in the one cell. The only limits are the number of species available and the range of dyes used. For example, a mouse derived primary antibody against protein A could be used alongside a goat derived primary antibody against protein B, so long as the anti-mouse and anti-goat secondary antibodies had different coloured labels.
 
The general protocol for antibody treatments is to expose the cells to the primary antibody for 1-2 hours, wash thoroughly in PBS to remove all unbound primary antibody, then expose to the secondary antibody for 1 hour. The cells are washed in PBS to removed any unbound secondary antibody, and washed in water to remove the PBS (which forms beautiful, but annoying crystals when the coverslips are mounted). To make a permanent preparation, the coverslips are mounted on a microscope slide in a special mounting medium which hardens upon contact with air.

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Targets

In this workshop, you will be demonstrating three components of the cells :

• The Nucleus – the HeLa H2B GFP cell line expresses a green fluorescent protein tag on the H2 Histone protein located in the nucleus. This protein is found throughout the nucleus and remains associated with the DNA, even during mitosis. These cells therefore show a green fluorescence in the nucleus, and in the chromosomes during mitosis.

• α-Tubulin (Alpha Tubulin) – α-Tubulin is one of the subunits of the tubulin protein which forms an important part of the cytoskeleton which provides support to the cell. During interphase, tubulin is found throughout the cytoplasm of the cell right up to the cell membrane and has a fibrous appearance. During mitosis, the cell assembles the mitotic spindle from Tubulin, and this structure appears as a “birdcage”-like structure on which the chromosomes are arranged. Tubulin is also a major component of the centrioles which make up the centrosome, and this appears as paired pinpoint bodies within the cytoplasm of the cell. In this investigation, the antibody against α-Tubulin used will be derived from a rabbit, and a secondary anti-rabbit antibody conjugated to a far-red (647nm) fluorophore will be used to demonstrate it. The human eye cannot detect light at 647nm easily, so a false colour (eg. magenta) will be applied, and the α-Tubulin will appear as fine fibres of this colour throughout the cytoplasm. Mitotic spindles will also be demonstrated using this method.

• Complex IV (or Cytochrome C Oxidase) – Complex IV is a large transmembrane protein composed of 13 subunits and found in the mitochondria of eukaryotic cells. Complex IV catalyses the last step in the electron transport chain in cellular respiration, transferring electrons to molecular oxygen and allowing the production of water molecules. In eukaryotic cells, apart from a couple of exceptions, Complex IV is found only in the membranes of mitochondria, which means that this protein complex can be used as a mitochondrial “marker” – a target which demonstrates the presence of mitochondria in the cell. The antibody against Complex IV used in this workshop is derived from a mouse, and a secondary anti-mouse antibody conjugated to a blue (405nm) fluorescent label will be used to demonstrate it. The mitochondria should appear as blue specks throughout the cytoplasm of the cell.

Fluorescence microscopy

In routine light microscopy, visible light is passed through a specimen on a microscope slide. This light continues up through the optics of the microscope to the eyes. Structures in a specimen will interfere slightly with the passage of the light, absorbing some wavelengths to give it a colour, or subtly bending it to reveal the shape of the structure. The structures visible through light microscopy may be enhanced through the use of coloured dyes which bind to components in the specimen, which is the basis of histochemistry, a useful technique to visualize elements in normally colourless animal tissue.

In fluorescence microscopy, structures are visualized using fluorescent dyes as labels. Fluoresence is a phenomenon where the chemical structure of the dye captures electromagnetic radiation of one wavelength (the excitation wavelength) and releases it as radiation of another, lower energy wavelength (the emission wavelength). For example, when light at 488nm (in the blue region of the visible spectrum) falls upon green fluorescent protein, electrons in the outer orbital of the atoms within the protein are excited to a higher energy state. When they return to their normal energy state, they emit photons of light at 509nm, which is in the green region of the visible spectrum (see figure below).

In fluorescence microscopy, light of the desired excitation wavelength is shone onto the specimen, exciting the fluorescent labels. The light emitted is passed up through the optics of the microscope to the eyes or a light detector. To prevent interference from reflected excitation light entering the optics, a dichroic mirror is used, which allows the emitted light to pass through, but not the excitation light. Further sensitivity can be achieved using emission filters, which only allows light of the desired emission wavelength to pass through. When light is shone down through the entire specimen, this is known as epifluorescence. Fluorescence microscopes have a number of adjustable filters, so that a range of excitation and emission wavelengths can be selected (see figure below).

 

Due to the arrangement of excitation and emission filters, fluorescence microscopes can only image a single fluorophore at a time. Therefore, in specimens which have multiple fluorophores, each with different excitation and emission wavelengths, multiple images must be taken. The microscope is set up for each fluorophore and an image taken before it is set up for the next fluorophore. The collected images, called channels, are then combined to create the final image. The light emitted by the fluorophores is often very weak, or at a wavelength which is difficult to detect with the eyes (the 647nm fluorophore used in this investigation, for example, cannot be seen by the human eye). Therefore, the digital cameras used in fluorescence microscopes take the images in grey scale as full colour detectors are less sensitive. Before the channels are combined, each is assigned a false colour which corresponds to the colour of the light emitted (see figure below).

Retaining the three channels in the final image can be useful for scientists, as the grayscale images make it easier to see finer detail than the colour ones, and comparing separate channels makes it easier to detect different fluorophores co-localised or situated close to one another. As a result, when presenting their results in lectures or scientific papers, scientists often present the three channels as grayscale images, followed by the combined colour image (see figure below).

A limitation of using epifluorescence microscopy is the difficulty in determining whether a combined signal is the result of two fluorophores in contact with each other (co-localisation). For example, if a green flurophore and a red fluorophore appear close to each other in the combined image, they appear yellow. This could be due to the structures they are attached to being in close physical contact, an important consideration when using fluorescence microscopy to determine interactions between proteins. However cells are three dimensional objects with a reasonably large depth of field, so the two fluorophores may not actually be close to each other, but actually overlapping through the depth of the cell (see figure below).

One solution to this problem is the use of confocal microscopy. In this method, background fluorescence is limited by the use of a pinhole aperture (at the expense of signal intensity, resulting in longer exposure times). In addition, the excitation of the fluorophores is done by tightly focused lasers which scan through the specimen. Therefore, only a particular portion of the specimen is exposed to the excitation wavelength and a virtual section is created. If structures are co-localised, they will appear on the same section, whereas if they are distant but overlapping, only one will appear on the section. Another benefit to confocal microscopy is that sequential sections can be stacked on top of one another to create a three dimensional image (a “Z stack”). Due to the lack of background fluorescence, confocal images appear clearer than epifluorescence images (see figure below).