Health and safety
Much biological research relies on the use of chemicals and organisms. Some of these are an inherent health and safety hazard. For example, most pure chemicals are irritants in certain doses, while others such as organic solvents and fungicides can be straight-up lethal to humans even in small doses.
This is why labs are run with a clear health and safety framework in place. This varies from lab to lab, depending on the research being carried out, and can include security clearance access to certain parts of the lab, designated first aid officers, clothing and personal protective equipment (PPE) guidelines and supervision systems.
For example, lab coats and nitrile gloves are standard procedure in most labs, and in more chemistry-focused labs, protective goggles too.
Beyond the risks present to lab workers, some items used in labs can be dangerous to other organisms and the environment once leaked into the outside world. Some chemicals will kill aquatic life or other animals, even if safe for humans. Therefore, depending on the usage of these chemicals, some labs will have a building-wide implementation of waste disposal and filtering as well as chemical dilution practices to minimise this risk.
Commonly, waste is incinerated to remove the risk of these materials reaching public spaces and the natural environment, whether they are toxic chemicals or genetically modified bacteria.
Liquids and solutions
Liquids and solutions are prepared in the lab as nutrients for growing microorganisms, buffers to hold proteins in their correct folding, chemical solutions to undergo various reactions, and anything else one’s mind can think up. In the lab setting, even plain water needs to be pure, and pH is measured to one decimal point. Liquids can be handled down to each individual tenth of a microlitre, which is a 10,000,000th of a litre.
A common task when handling liquids in an experiment is the need to dilute a stock into multiple serial dilutions with decreasing concentration of the solute in the solvent i.e. glycerol in water. For example, 5 tests are undertaken, each a tenth of the concentration of the original solution. So a 50% glycerol solution of 10 mL (which contains 5 mL of glycerol and 5 mL of pure water) is used to create the subsequent 4 solutions.
In this serial dilution, a tenth (i.e. 1 mL) of the original solution is added to a new tube, and made up to 10 mL with 9 mL of pure water. This is now a 5% glycerol solution. A tenth of this is used to make the next solution of 0.5% glycerol, and so forth. This is a log dilution series because each dilution is a factor (in this case of 10) less concentrated than the previous.
Depending on how dilute we need the solution to become in our experiment (sometimes diluting too much can prevent e.g. a chemical reaction from occurring) we can also do a linear dilution series where each dilution is half of the previous one. In the 10 mL of 50% glycerol example, this amounts to aliquoting 5 mL of the solution into a new tube and adding the remainder 5 mL of pure water to make up 10 mL total volume. This creates a 25% glycerol solution, which is then used for the next dilution by taking out 5 mL (half) of it and adding to the next tube, making it up to 10 mL with another 5 mL of water. This is now the 12.5% glycerol solution, and so on and so forth.
The protein concentration in solution can be measured by its optical density. This means that how much light passes through the sample is correlated with how much solute there is in the solution. A special instrument called a colorimeter (or spectrophotometer) can work this out.
The data can be plotted with a standard curve. Since the data points have a linear relationship, all the unknown points on the same curve can be derived from the graph. For example, the point on the curve corresponding to an optical density of 0.3 is equivalent to a protein concentration of 0.22 mg/ml.
pH affects enzyme activity and chemicals involved in metabolism, so maintaining optimal pH is key to enabling the right outcomes from microorganism culture and various biochemical reactions. Buffers can be pH adjusted e.g. to pH 7.4 by using a pH meter, acid and alkali solutions. Commonly, sodium hydroxide (NaOH) is used as the alkali (providing the OH– groups), and hydrochloric acid (HCl) is used as the acid (providing the H+ ions).
If the measured solution is too low (on the pH scale of 1 -14; most acidic – most alkaline), pH can be increased by adding NaOH to make it more alkaline. If it’s too high, HCl can be added to make it more acidic. Buffers are able to withstand fluctuations in pH brought about by adding various chemicals to the solution, and bring the pH back to the pH set initially. Therefore, buffer solutions are a key foundation of biochemistry and molecular biology.
Often, solutions contain multiple types of component that need to be separated. Whether this is a bacterial culture that needs the bacteria separated from their liquid media, or a mixture of proteins from which we need to isolate just one kind, there are multiple approaches to achieving separation.
Centrifugation is one of the basic go-to methods of separating components based on their physical properties and interactions with each other – but chiefly mass. Simply put, heavier things are pulled in closer through gravity. Earth’s gravity doesn’t go far enough to separate many things. Centrifugation (spinning) is a way of increasing the g force on the components many times higher, so that this separation takes place.
The components with a higher mass accumulate to the bottom of the centrifugation tube, while the lighter things remain on top. A clear dividing line between components of different density can often be observed. The components at the bottom, often solids, are termed the pellet, while the components on top, often liquids, are termed the supernatant.
Depending on the experiment, either the pellet or the supernatant is the component of interest. The other component is often discarded. As always, knowing which is which is key!
In order to separate smaller components, like amino acids and proteins, highly specific methods can be employed, including affinity chromatography. This is more specific than the basic chromatography technique done to separate different chemicals from a mixture e.g. paper chromatography and thin layer chromatography.
Paper chromatography involves using a defined piece of chromatography paper and placing a droplet of the mixture at the bottom, in the middle of the paper. This section is then immersed in a solvent which is drawn up the section of paper through capillary action. Depending on their chemical properties, some components of the mixture will be drawn up with the solvent, while others will lag behind or not move at all. This separation is enabled by their interaction with the stationary phase (the paper) and the mobile phase (the solvent).
Thin layer chromatography uses the same concept, except it uses an aluminium, glass or plastic layer instead of chromatography paper. It is used more commonly, as paper chromatography has become more of a teaching tool than research tool. Thin layer chromatography can be used in organic chemistry research to indicate whether a starting compound used in reaction has been used up or is still present.
Affinity chromatography is the specific type of chromatography used in molecular biology research, as it enables the separation of given proteins from mixtures of bacterial cell lysates, buffers, etc. The concept of affinity is that some specific property of the target protein (size, presence of tags previously added for this purpose, chemical bonding including hydrogen bonding, disulfide bridges, ionic interactions, etc.) can be used to bind it to a stationary phase, while compounds that do not meet that criteria can be washed away. The protein can then be eluted into a solution that contains the target protein only, resulting in its purification from the original mixture.
In order to preserve the chemical environment for each step, different buffers are used for loading, washing (separating) and eluting. A common binding tag used to separate proteins is a His tag which consists of 6 histidine amino acids joined together. It can be genetically engineered into any DNA that codes for the target protein. Therefore, the protein will be synthesised with an extra part – the His tag – at the start or end of its sequence. The His tag can then be used to bind the target protein to the stationary phase, and separate it from all the other proteins in the mixture (without the tag) using a nickel column.
You can see it’s a nickel column just by its blue colour. Purification columns for chromatography come in many shapes and sizes. The smaller ones can be used for small volumes, while the largest ones are used in conjunction with automatic pumping machines.
Manual use involves passing the original mixed solution with the target protein through the column (loading), followed by washing the column multiple times with a special buffer to remove the unwanted components. Lastly, and the most critical step, is elution. This washes away the bound target protein from the column in a yet another specialised buffer, where it can be stored for longer in its purified medium.
Alternatively, all the buffers can be pre-made and used with an automated machine. The machine does all the buffer changing and pumping, and it can even collect fractions of the solutions obtained. Only some of these fractions will contain the target protein.
Protein electrophoresis is a method for separating smaller amounts of protein mixtures for the purpose of visualising them by size and charge. The principle is the same as DNA electrophoresis. The mixture is loaded into a porous gel, and a current is passed through it. The proteins travel down the gel, away from the negative end of the current, towards the positive end, as they themselves have a negative charge.
The gel is removed from the tank and stained with a dye to highlight the protein bands. A marker, or ladder, is used adjacent to the tested protein samples in order to provide a reference for the sizes on the gel. As you can see, the smaller fragments travel faster than the larger fragments. This is because smaller fragments can make their way more easily through the gel matrix than larger fragments.
Protein size is measured in Daltons (more specifically, kilodaltons). For example, human insulin is 11.98 kDa. On the gel, it should show up just above the 11 kDa marker, in a parallel lane. Each parallel lane is a separate protein sample.
If we are to separate proteins because they are a contaminant rather than a product of interest, e.g. casein in cheesemaking, protein precipitation at its isoelectric point can be carried out. A protein’s isoelectric point is the pH in which its positive and negative charges on its amino acid residues even out, making the protein neutral.
Precipitation (coming out of solution) is the attraction between proteins that brings them closer, resulting in their aggregation out of the solution they are in. Mineral acids such as hydrochloric acid are used to achieve isolectric point (pI) protein precipitation. The isoelectric point of proteins differs between them, with most being around pH 4-6.
Bringing the pH of the solution to their isoelectric point causes precipitation because it removes repulsive forces (at a pH higher/lower than pI, the surface of proteins is predominantly negatively/positively charged, causing them to repel each other), while allowing the attraction forces between proteins. This causes precipitation out of solution.
Due to the irreversible denaturing (protein misfolding that compromises its function) effects of the acids used to cause precipitation, isoelectric point precipitation cannot be carried out on proteins of interest that must be kept functional. Therefore, it is only used to get rid of contaminant proteins to be discarded.
The high specificity of the antigen-antibody bond makes use of antibodies as a lab technique high on the list. Antibodies are used with proteins as a labelling method to detect presence of the target moiety (in biochemical reactions, testing or disease detection), or with whole tissue samples to detect presence of specific organelles for visualisation under a microscope.
Assays are carried out in multi-well plates to detect the presence of a specific antigen, against which an antibody is added. If the antibody binds to the antigen, it causes a shift in a reporter enzyme added or linked to it, resulting in a colour change quantifiable by a spectrophotometer.
Antigens are any identifiable, specific parts of a molecule, and can be found on proteins, tissue samples, patient samples, pathogens, etc. Antibodies recognise antigens specific to them. The antigen-antibody relationship is at the heart of an organism’s self-identification and identification of invading species.
The enzyme-linked immunosorbent assay (ELISA) is a common immunoassay used to detect antigens in samples. It relies on adding the samples to a multi-well plate made of polystyrene, that can immobilise the antigen-containing molecules. The specific antibody is then added and allowed to bind, if it is to bind. After washing away the unbound molecules, the reporter enzyme is added to produce the colour change. Voila!
There are other versions of ELISA where the “capture” antibody is immobilised to the plate well instead of the sample, and the sample is added on top. The key principles of antigen-antibody binding, and antibody-indicator binding/activating are the same.
The antibody for the antigen is the primary antibody, while the “second antibody” binding the primary antibody (if applicable) is the secondary antibody. The latter can be the one with the indicator attached that produces a colour change.
The principle of primary antibody and secondary antibody is key to fluorescent labelling. The primary antibody selects for the target antigen, while the secondary antibody selects for the primary antibody (now bound to the target antigen; unbound antibodies are washed away) and provides the colour change. Since this labelling is sometimes done on a very small scale, the colour change is measured by fluorescence using a special detector that displays the product through a filter, or a microscope.
Western blotting is a common technique that involves visualising very small amounts of protein on a gel (following electrophoresis, see previously) by using a highly specific primary antibody. It’s possible to have this primary antibody produce the fluorescence itself, in which case a secondary antibody isn’t necessary. Either way, the Western blot is a very specific method for detecting proteins. The proteins are transferred to a nitrocellulose paper from the gel by sandwiching them under a current before the antibody treatment is carried out.
Whole tissue samples can be labelled fluorescent using antibodies and visualised with a microscope. Specific antibodies can target DNA, nuclei, muscle fibres, etc. and produce different fluorescence colours under a microscope, to help differentiate them.
This is where secondary antibodies come in handy. Each with a different colour signal, they can be selected to be different for different cell organelles, to help see them clearly and as separate.
Making monoclonal antibodies
In order to manufacture all these types of highly specific antibodies (produced in vivo by B lymphocytes), identical clone cells of each specific B lymphocyte are cultured and selected for. The cells that originate from one parent cell, and are therefore identical to each other, are termed monoclonal. Hence, their antibody products are monoclonal antibodies.
The neat trick to achieve this is by hybridising extracted B lymphocytes (e.g. from an animal that has been exposed to a certain antigen and therefore has mounted an antibody response against it) with their counterpart cancerous B lymphocytes (myeloma). The resulting cells, called hybridomas, can propagate by division because they are cancerous, while maintaining the ability to produce antibodies.
A common lab chemical called polyethylene glycol (PEG) can be used to create the fusion between plasma membranes (hybridisation), although a more successful method is using a selective growth medium that only allows hybridised cells to replicate. This naturally selecting environment results in easy hybridoma culture, as non-hybridised cells simply die away.
Single cells from this hybridoma culture can then be used to start fresh cultures to grow specific antibodies.
You will need to know about the difference between bright-field (visible light), transmission electron and scanning electron microscopes – LM, TEM and SEM. Both the latter (as the name suggests) use a beam of electrons, rather than light, to produce an image of the sample.
TEM uses electrons which pass through the sample, so the resulting micrograph (image) shows everything within the sample in black and white, for example organelles in a cell. SEM uses electrons which scan the sample in 3D, resulting in a coloured micrograph with 3D detail, but no components from within the sample.
In bright-field (light) microscopy, light does go through the sample, but the outcome depends on the thickness of the sample. For example, the plant root slice in the diagram (LM) is thin enough to be able to see through the thickness of the sample. Light would also travel freely through air but not various materials of high opacity.
When talking about microscopes, differentiating between resolution and magnification is important. In principle, it’s not hard to understand. Imagine zooming in a photo to try to see a detail. That is magnification. Now imagine the photo has a low resolution, and if you magnify it, you can only see annoying pixels. If the image had a high resolution, you would be able to see the detail clearly after zooming in. So magnifying is zooming in, while resolution is the focus power. You will need to be able to calculate actual sizes and magnifications of various drawings. The equation for that is Image size on paper = Magnification x Actual size. This gives magnification = image size on paper / actual size. “I AM” summarises it nicely in a triangle.
Staining is a key precursor to microscopy. Most samples would not register well under a microscope without some form of staining. This can also be critical to the experiment carried out. For example, we might need a stain for the cell nucleus as well as a stain for the cell fibres.
For microscopes with fluorescent wavelength filters, fluorescent stains are used. These can be bound to very specific antibodies to target specific cell types or cell organelles, as previously mentioned.
Muscle cells can be stained for their nuclei (blue) or one of the constituent proteins like actin (red). Stains of different colours and hence wavelengths are used to differentiate the various parts that we want to visualise. Advanced microscopes such as confocal microscopes that use lasers can focus on multiple points in the sample, and relay multiple wavelengths at the same time to create an impression of a section through the sample, add together the data through the multiple layers in the sample, and create complex images of specimens.
Aseptic technique and cell culture
In a similar line of thought with the aseptic techniques employed in microbacterial culture (disinfection of work area and tools with IMS, using hoods for air flow control, etc.), aseptic techniques used for culturing cells from large eukaryotic organisms such as mammals are an essential backdrop to working with them.
Infection of cultured cells can lead to their death. When as long as many months could have been spent growing cells e.g. brain cells, having all that work be wiped out in an instant by a bacterial contamination is clearly to be avoided. If research relies on deep-frozen samples of cells that themselves have been contaminated during the conservation process, then potentially a whole batch of cells could have been compromised.
Therefore, fundamental steps are taken to protect cells used for culturing. Cells are cultured in specially shaped, horizontal flasks. Their neck and lid have been designed to minimise the risk of contamination. Whenever opened, it must be done in a sterile hood (called a laminar flow cabinet) that has been pre-disinfected with 70% IMS (industrial methylated spirit, previously ethanol).
The hood maintains an air flow towards the outside to prevent microorganisms travelling through the air entering the working area inside the hood. Note that this is the opposite of a chemistry fume hood which maintains air flow towards the inside and out of the lab to protect the worker from toxic vapours.
Despite superficial impressions of cultured cells (often stem cells), that they are a limitless supply of life, cells are just delimited volumes of space that reflect their immediate environment. As such, they rely on highly specialised media to feed on and develop. Since chemical manufacturing of biological compounds such as proteins is not advanced enough to enable easy production of these media, the most convenient route to obtaining them is as a byproduct of the meat industry. Examples of media include animal blood and isolated fluids from different tissues.
Serum from blood contains many growth factors that stimulate cell development, as well as specific molecules that can determine cell differentiation. These can be species-specific, so they have to be sourced from the right species for each cell line.
The source of these cells includes explants of tissue from organisms (tissue samples removed from the body) or a few starting cells from previous cell line cultures, some of which are commercially available.
Once again, similarly to bacterial cell culture, these cells can be quantified by using a special marking device called a haemocytometer under a microscope. A very small volume of the cell culture (following unsticking them from the culture flask using the enzyme trypsin) is pipetted under a glass slide over a marked area (the grid), and cells in a small section of the grid are counted to work out the total.
For example, if there were 9 cells in the 10x objective square visible from the grid, it could be multiplied by 16 to obtain the total number of cells found in the top right-hand square of the grid, and again by 9 to obtain the total of the whole grid. Then, the total could be further multiplied to get the total cells in the whole cultured volume. If this was a 10 microlitre cell culture sample used in the haemocytometer, we would multiply the total number of cells by, e.g. 1,500 to get the total number of cells for a 15,000 microlitre (15 ml) cell culture.
In this case, that would be 9 x 16 x 9 x 1,500 = 1,944,000 cells in the 15 ml culture. This could be expressed as (1,944,000 / 15 = 129,600) 129,600 cells per ml. An even further shorthand for this is 129,600 cells ml-1.
Growing different cell lines differs based on the cell type. Primary cell lines (derived directly from source e.g. patient tissue) cannot usually be cultured indefinitely. After an initial period of growth (several days to weeks), they begin undergoing senescence and die.
Immortal cell lines, such as those isolated from cancer, have different properties and can grow indefinitely. This makes them useful for research, although their mutated state may interfere with experimentation and comparison to non-mutated cells. They can be preserved under very cold, liquid nitrogen storage facilities and used later.
For plant tissue culture, a similar aseptic approach is taken by preparing plant tissue in laminar flow cabinets. Explants (samples) are commonly grown to produce plant clones, as they can reproduce asexually from many different parts of a whole plant. This practice is called micropropagation.
Plant tissues also respond acutely to environmental stimuli, notably growth regulators such as the plant hormones auxin and cytokinin. Excess auxin would lead to too much root development, while excess cytokinin would lead to too much shoot development. Their control is therefore key to directing adequate plant growth.