General Notes

This chapter explains a number of ideas that need to be understood in order to fully understand the book. There are notes and discussion regarding chemistry, nomenclature, and general lab etiquette. Additionally, there are subsections on aseptic technique and aliquoting, both of which will benefit someone new to the laboratory setting. Some of the ideas explained in this chapter are ideas that are important but did not fit well into one specific technique—since they were necessary, they are included with general notes. This chapter assumes that the reader understands pH, molarity, and molality, as well as how to calculate solution molarities from gram weights and solution volumes. The Aseptic Technique section describes common methods to avoid introducing bacteria or mold into experiments, while Aliquoting describes what aliquoting is and how to apply it to the reader’s own experiments and setup.

Keywords

Aseptic; aliquot; solutions; pipetting; multiplexing; troubleshooting; etiquette

Basic molecular protocols require a basic understanding of solutions chemistry (i.e., the concepts of molarity, molality, pH, and stoichiometry). If you do not know how to calculate molarity from weight (in grams) and volume, or how to calculate the grams you need for your solution from a molarity value, then please review a basic chemistry text. Otherwise, some of these instructions will be incomprehensible.

Typically, solutions are stored and labeled at some molarity concentration (M, mM, µM, etc.), but many protocols and recipes refer to a multiplier value, such as 10×, 5×, or 1×. The “×” is a multiplier and tells you how much you’ll need to dilute that stock solution. Unless the protocol says otherwise, you will generally use solutions at a 1× concentration. For example, if I have a 10× stock solution and I want 1 liter of 1× working solution, I will dilute 100 milliliters (ml) of 10× stock with 900 ml of water to make my 1× working solution (i.e., divide a liter into 10 parts to find the amount of stock solution to use).

Remember your metric prefixes. Using molarity (M) as an example: millimolar is 10−3 molar (mM), micromolar is 10−6 molar (µM), nanomolar is 10−9 molar (nM), and picomolar is 10−12 molar (pM).

“Multiplexing” means that you run multiple experiments in a single tube or tissue sample (or similar reaction site). Multiplexing a quantitative polymerase chain reaction (PCR) (see Chapter 4) means that, using unique probes for each target, you amplify multiple targets within a single tube or well. In immunoblotting and immunohistochemistry, multiplexing would mean that you use two or more antibodies on a single blot/tissue slice at the same time and visualize them at the same time (possible via fluorescence and some colorimetric reactions). In in situ hybridization, multiplexing means using two or more probes in the tissue at the same time and visualize them at the same time (again, via fluorescence and some colorimetric reactions). Multiplex reactions are great if you can get them, but be aware that the optimal conditions for one reaction may be the worst conditions for the other reaction. Additionally, if you have a limited amount of one crucial reagent, such as deoxynucleotide triphosphates (dNTPs), that limiting reagent will be used for all of your reactions simultaneously and each reaction will, therefore, affect the others.

“Vortexing” means that you use a device called a vortexer (we scientists are a creative lot) until you see the “cyclone” in the center of the tube. This can rapidly mix solutions, but it is inappropriate if your solution components are sensitive to mechanical forces. For example, DNA is sensitive to mechanical force and solutions containing DNA should not be mixed via vortexer.

“Pipetting” means that you are using pipets to measure some volume of whatever solutions you are using in your experiments. Pipetting is a broad term, encompassing the use of rubber balls on the ends of labeled pipets, micropipettors and disposable tips, disposable bulb pipets, and labeled pipets with hand-held electrical pumps (called pipette guns). When in doubt, pipette solutions in and out slowly and make sure the liquid is all out or nearly so. When changing the volume measurement on the micropipettor, perform it exactly how the manufacturer stipulates—if they have a wheel in the middle, then using the top will eventually take the top off! If they don’t have a wheel, then turn the top of the micropipettor.

Develop your pipetting technique. Many of these experiments depend on your ability to accurately pipette the correct amount of fluid. You should pipette solutions slowly and evenly for best results. After much practice, you will be able to quickly pipette accurate amounts of solution. Additionally, it is common and necessary to mix solutions by pipetting the fluid in and out of the tip repeatedly. Nucleotides, among other things, like to stick to plastic, and pipetting back and forth brings them back into solution. If you did not mix thoroughly, the amount of liquid you pipette might be correct, but the things inside the liquid that you want may not be at the correct concentration. I prefer to count my passes (one pass=into the pipette tip and back into the container), and consider 30 passes to be mixed sufficiently. Finally, plan your pipetting to a minimum number of steps. Every time you pipet, you increase the risk of experimenter-based errors and contamination. For example, if I plan to pipette small volumes three times, I have fewer chances to contaminate my solutions than if I pipette smaller volumes six times. Even better, use a different micropipettor and pipette once.

When adjusting the pH of Tris-based solutions, use a Tris electrode-based pH meter. When adjusting the pH of paraformaldehyde (PFA) solutions, use disposable pH strips (most pH meter electrodes aren’t tested against PFA solutions, so they may damage the electrode). For other solutions, a standard pH meter is fine. Always continuously stir solutions (either by hand or by stir bar) while adjusting the pH, or else you may measure the pH of your acid or base instead of the solution as a whole. Follow manufacturer instructions for proper storage of your pH electrode. If you do not know how to use pH solutions, the manufacturers of pH meters have instructions included with their products or on their web sites. Additionally, you should be able to ask someone in the lab if those instructions don’t work.

A freeze-thaw cycle is, any time where a frozen solution is thawed, used, then frozen again. Just about anything that you want to freeze is sensitive to the number of times it is thawed. This is the big reason that you should use aliquot solutions (see Aliquoting section) and avoid frost-free freezers. Frost-free freezers avoid forming frost by essentially thawing at some critical point (a specific temperature, time, or other factor), which means that your samples in a frost-free freezer will go through many freeze-thaw cycles without you knowing it. Repeated freeze-thaw cycles will render your antibodies useless, degrade guanosine-5′-triphosphate (GTP), and degrade nucleotides.

When troubleshooting, a “known good” has immense value. A “known good” is something that you know works: a wire that you’ve tested and conducts electricity as it should, an antibody that gives great results every time, a PCR primer set that is specific, or a selection of tissue that has generated great results in the past. Being able to test a known good against your new, unknown results can eliminate a number of variables from your troubleshooting (if your known good antibody doesn’t generate staining, for example, then the problem is not necessarily the antibody but could be the signal-generating components or your sample preparation).

“Common use equipment” is typically a euphemism for “no one takes care of this machine.” Even if someone does take care of the equipment (you should be grateful if this is the case), it will still benefit you greatly to read the user’s manual and, if necessary, perform routine maintenance days or hours before you use the machine (this is a must if no one takes care of the machine). By performing maintenance before your experiment, you lower the risk of having the machine break down at a crucial point and ruining your experiments. Finally, clean up after yourself when using a common-use machine. No one needs to examine a spill in a common centrifuge and wonder if that spill is safe or contains a deadly virus.

Some abbreviations are used in multiple places and mean different things. You may find a lab notebook that refers to RT as reverse transcription, room temperature, or real time. While I will avoid that in this book, always look for context clues as to which abbreviation means what.

If you are borrowing equipment or space from anyone else, remember: leave a place better than you found it. If you happen upon a lab with impeccably clean and organized shelves and perfect conditions, then do two things. First, take a picture for posterity, and second, leave the lab in the same impeccable condition that you found it.

Aseptic Technique

Before starting any of these molecular biology experiments, it is important to understand and be willing to refine your aseptic technique. As I understand it, “aseptic technique” is a phrase from microbiology but is perfectly applicable to molecular biology. Proper aseptic technique avoids introducing any...