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

John T. Corthell

• Publisher: Academic Press
• Year: 2014
• ISBN:
  9780128015278
• Topic: Biological Sciences
• Subtopic: Neuroscience
• Index: Biological Sciences

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

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 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 unwanted materials (bacteria, molds, or unwanted solutions) into your reactions.

A very important factor in aseptic technique is sterilizing solutions. There are many ways to sterilize solutions, but only a few are important in practice: autoclaving, filter sterilization, and bleach. Bleach is used when you have a dangerous material to dispose off and you need to be absolutely certain that material is absolutely dead before it goes into the trash (where your unsuspecting janitorial staff could be hurt if you are negligent). Autoclaving is using a combination of heat and pressure to kill bacteria and destroy bacterial endospores. Autoclaving takes place in a specially designed machine called (ta-da!) an autoclave. Autoclaves have multiple settings and can vary by manufacturer. One uses autoclave tape to seal or mark items for autoclaving; autoclave tape is designed to have specific patterns after autoclaving to signal that your item is sterilized. In general, 5–20 min of autoclaving is enough for plastic tips and tubes (check the melting temperature by manufacturer to make sure your plastics won’t melt in the autoclave). Autoclave-safe bottles need to have lids on loosely. Try not to mix biohazardous waste with nonbiohazardous materials in the autoclave, as anything dangerous needs more time in the autoclave for people to be absolutely certain that it is no longer a threat. Filter sterilization is for solutions that contain vitamins or components that break down due to the heat and pressure of autoclaving. Check every component of your solutions to make sure that you can autoclave them and still have the intended solution.

Other than starting with clean solutions, using a great deal of proper aseptic technique is common sense. If your pipette tip just fell onto the lab bench, you do not use that tip for your experiment; you throw it away immediately. If you do use that tip, you won’t know if the experiment failed because that’s the true result, or if it was because you introduced bacteria that treated your samples as food. If the experiment works, was it because your experiment was a success or did you get a false positive from that mold you inadvertently introduced? I knew someone who thought for 6 months that they had discovered a completely new protein in their samples (“I’ll be famous!”), only to discover that they contaminated their samples with their own skin cells for 6 months. Developing your aseptic technique is very important to avoid wasting your time and funds. Here are some ideas to get you started:

1. Wear gloves. Always wear disposable gloves and relevant personal protective equipment. You wouldn’t believe how many people don’t use gloves and wonder why their samples look funky (it’s a technical term). Remember that “molecular biology” means “wear gloves.” You wear gloves in some cases to protect yourself, but here you wear gloves to protect your samples from you and your bacteria. If you are working with pathogenic bacteria or viruses, then you are protecting yourself and protecting your samples. Remember that working without gloves while experimenting on dangerous materials (Ebola virus, HIV) will only hurt you. If necessary, gloves can be autoclaved in commercially available autoclave bags or wrapped in paper towels then aluminum foil and sealed with autoclave tape.

2. Only use autoclaved pipette tips. Some manufacturers sell RNase-, DNase-, and pyrogenase-free tips. These tips are fine to use; you just want to make sure that your tips have been sterilized or cleaned up in some way (aerosol-free tips help with pipetting but don’t affect aseptic properties). Autoclaving doesn’t necessarily get rid of RNases (nucleases that destroy RNA and may therefore destroy your samples), but the plastics can’t handle the multiple hours at 200°C that are guaranteed to destroy RNases. In practice, autoclaving works just fine for tips and plastic sample tubes. Tubes and tips that are in large bags are exposed to bacteria and other things as soon as you open the bag, so if you purchase those items in bulk bags, autoclave them in separate containers to ensure their cleanliness. If you drop your tips on the floor or a lab bench, don’t use them. If you are really cheap, they can be washed and reautoclaved (people do this), but don’t ruin your experiment by using dirty tips. A tip costs less than 10 cents (USD); what is that compared to the cost of the rest of your experiment?

3. Autoclave or otherwise clean your tools as necessary. It should go without saying that any tools used for surgery or dissection need to be washed after use. Tools can be autoclaved in the same manner as gloves, though scissors and other sharp implements may require resharpening after autoclaving. Tools can also be washed with 70% ethanol (this percentage works best for killing bacteria, compared to 100% ethanol or 50% ethanol solutions) and/or commercially available RNase-destroying solutions. Some people dip their tools directly into RNase inhibitors between samples.

4. Autoclave or otherwise sterilize all solutions that you will use for PCR. Most solutions are sold as sterile or cleaned solutions; these are fine unless you have some reason not to trust them. However, if you do not trust the solution or you made it from scratch and used nonsterile components, sterilize the solutions. Sterile solutions are a necessity for PCR, electrophysiology, immunoblotting, immunohistochemistry, and for long-term sample storage (check Solutions in those sections to see about which solutions are sterilized and which are not).

5. Clean up the working area before each use. If you can, designate a single area for each type of experiment (a PCR setup area, separate from where your reaction occurs). If you don’t have a designated area for performing these experiments, clean up the bench with 70% ethanol. If you are working with RNA, you can also use RNase-destroying solutions to clean your workspace.

6. Perform work in a “known good” clean area. This relates to the previous tip. If you can, having a designated space that is kept clean (a dedicated sterile hood, for example) can be quite beneficial. The sterile hood keeps particles out via HEPA filters, while you keep the bottom clean by washing with 70% ethanol before and after each use. I keep sterilized pipettors, pipette tips, and tubes in the dedicated hood to avoid introducing new contaminants on a regular basis. Common-use tips, tubes, and pipettors are a small price to pay for people not introducing new molds, bacteria, and junk into your sterile hood by bringing their equipment with them every time they want to run an experiment.

7. If your tip touched something that is not sterile, throw it out and don’t introduce it to your experimental solution. Let’s say you grabbed a capped tube with your bare hands and then put on gloves. The inside of the tube should still be sterile, and so you can use it without worries. However, while pipetting, the end of your tip touches the outside of the tube. Throw out the tip and start over. You can’t trust the tip to not introduce skin cells, skin oils, skin bacteria, or other junk into the reaction. Don’t sabotage a $200 experiment for the sake of $0.07 of tips. See number 2, above.

You will find that, for different experiments, there may be additional steps you take to keep your samples clean. These suggestions are just to get you started.

Aliquoting

Anything that is sensitive to freeze-thaw cycles but needs to be stored in your freezer should be aliquoted. Aliquoting is the practice of separating solutions into smaller batches in separate tubes. The idea is that if you ...

Table of contents

• Cover image
• Title page
• Table of Contents
• Copyright
• Introduction
• Chapter 1. General Notes
• Chapter 2. DNA and RNA Extraction Protocols
• Chapter 3. Agarose Gel Electrophoresis
• Chapter 4. Reverse Transcription (RT) and Polymerase Chain Reaction (PCR)
• Chapter 5. Antibodies and Titrations
• Chapter 6. Protein Extraction
• Chapter 7. Immunoblotting (Western Blot)
• Chapter 8. Immunoprecipitation
• Chapter 9. Perfusion and Immersion Fixation
• Chapter 10. Immunohistochemistry
• Chapter 11. In Situ Hybridization
• References