Basic Molecular Protocols in Neuroscience: Tips, Tricks, and Pitfalls

By John T. Corthell

Basic Neuroscience Protocols: Tips, Tricks, and Pitfalls contains explanatory sections that describe the techniques and what each technique really tells the researcher on a scientific level. These explanations describe relevant controls, troubleshooting, and reaction components for some of the most widely used neuroscience protocols that remain difficult for many neuroscientists to implement successfully. Having this additional information will help researchers ensure that their experiments work the first time, and will also minimize the time spent working on a technique only to discover that the problem was them, and not their materials.

• Describes techniques in very specific detail with step-by-step instructions, giving researchers in-depth understanding
• Offers many details not present in other protocol books
• Describes relevant controls for each technique and what those controls mean
• Chapters include references (key articles, books, protocols) for additional study
• Describes both the techniques and the habits necessary to get quality results, such as aseptic technique, aliquoting, and general laboratory rules

• Language: English
• Publisher: Academic Press
• Release date: Apr 16, 2014
• ISBN: 9780128015278

John T. Corthell

Dr. Corthell received a PhD in Neuroscience from Florida State University, where he studied circadian rhythms using whole-cell patch-clamp electrophysiology, as well as numerous molecular biology techniques, in the olfactory system. He has more than 10 years of experience in laboratory work and techniques in the biological and chemical sciences, and more than 5 years teaching. The author has cultivated these techniques over the years through reading, asking questions, and trial and error. Each of these techniques has been tested, performed, and in some cases optimized by the author himself. He enjoys reading, writing, learning, children, and big dogs. When not working, he longs to go SCUBA diving.

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Chapter 1

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