General guidelines for molecular biology labs

There are several basic molecular techniques that are not mentioned specifically in molecular biology protocols. Please review this guide before every lab until the techniques become second nature.

Following Protocols

It is crucial that protocols are not only followed carefully, but that you understand the purpose of each step. Frequently complications will be encountered and you can effectively trouble shoot only if you understand how the protocol works.

Since we are attempting to give you a real research experience, the protocols are presented as they would be in a research lab. This means the protocols are written much more concisely than you may be used to. For example, a protocol will not include details like “label your tubes” as it is generally understood that unlabelled tubes will lead to disaster.

When following a protocol, you must think and anticipate.

Using a protocol effectively & efficiently

  1. Read the entire protocol ahead of time, making a list of any equipment or components needed. Make sure you understand what you need to do in each step – look up words you do not understand.
  2. Check that you have all the components and equipment needed and that you know how to use them.
  3. As you work through the protocol, read each step in its entirety, right before you do it.
  4. Anticipate. For example, if you will need another set of labelled tubes in step 5, prepare these tubes during a 5 min spin in step 3.


Biological molecules can be damaged or degraded by enzymes that are found on non-sterile surfaces. Consequently, all manipulations are done with sterile materials (e.g. sterile tips and tubes). In our lab, most consumables and solutions will be sterilized by autoclaving. Sterile tube and tip packages have autoclave tape attached; this tape has dark stripes if the materials have been autoclaved.

To protect the sterility of your materials

  • clean your bench with the provided 70% ethanol at the beginning and end of every lab period;
  • keep containers closed, except when removing the contents (i.e. do not leave the tip box open);
  • do not touch the materials with your hands or any other non-sterile surfaces.


We will frequently work with living organisms, many of which have modified DNA and; therefore, are classified as Genetically Modified Organisms (GMOs). GMOs are considered biohazards until their potential effects on humans and the environment are assessed (this is rarely done for products produced for research purposes). DNA and proteins extracted from such organisms are also classified as biohazards. The organisms we are working with have been used by 1000s of researchers for decades with no evidence of ill effects, none-the-less it is appropriate and required by law that we take certain precautions.

  • GMO cultures and any materials derived from GMOs must be destroyed before disposal. The materials we use are destroyed using high-pressure steam. Your job is to make sure all materials you use – tips, tubes, gloves etc. are placed in the bio-hazardous waste buckets – I will take care of the waste from there.
  • Rather than trying to track which tubes came in contact with the bio-hazardous material – assume everything you use is bio-hazardous and treat accordingly.
  • Please report any spills to your lab instructor immediately – you will not be in trouble – we simply need to help you clean up.
  • Always wash your hands thoroughly before leaving the lab with soap and hot water.


General lab safety is discussed here but it is worth repeating a few points specific to this lab. Besides the biohazards mentioned above please be aware of the following hazards in the lab:

  • Bunsen burners – tie back long hair and avoid loose clothing.
  • Bunsen burners and ethanol – ethanol is very flammable and we often use it with an open flame. Do not panic if you accidentally set ethanol on fire. If the ethanol is in a container put the lid on the container, if the ethanol fire is on the bench and is small – turn off the burner and move away any flammable objects – the fire will quickly burn itself out. If the fire seems large, notify your instructor immediately.
  • We will occasionally use hazardous chemicals such as ethidium bromide – please wear gloves and eye protection when instructed to do so and always wash your hands before leaving the lab.
  • We regularly use equipment such as centrifuges and vortexes that can be hazardous when used incorrectly – make sure you are using all equipment correctly and be aware of what is going on around you.
  • Watch out for heat blocks that have very hot surfaces.

Proper use of equipment and reagents


Please review Proper use and care of pipetters. Your pipetting skills will be tested in the first lab. Remember to keep tips sterile (see above).


  • Centrifuges must always be balanced before being run. This means that every tube must be directly across from another tube that has an equal mass.
  • When using a low speed micro centrifuge (max rpm ≦ 15000) you can assume that tubes with identical contents and volumes have the same mass (e.g. if you are doing two plasmid preps side by side the tubes should have the same components and volumes).
  • If the tubes do not have identical components or do not have the same volume you will have to weigh the tubes and adjust the lower weight tube to be the same weight as the heavier tube (+/- 0.1 g).
  • If you can't adjust the volume of the tube, or if you only have one tube to spin, make up a similar mass tube using water (we call this a balance-tube).

Once the centrifuge is loaded

  1. place the rotor lid on the rotor
  2. close the centrifuge lid
  3. set the speed (make sure you know what the speed should be)
  4. set the timer (setting the timer usually starts the centrifuge)

Always place your tubes with the hinge facing the outer rim of the rotor.
If you are spinning something down this will make it easier to find the pellet.

Working with reagents

When repeatedly pipetting the same reagent into multiple tubes, you must change tips between tubes. This prevents contaminating the reagent or cross contamination between samples (one exception – if the tube is empty you can use one tip to add the reagent to all tubes).

A note on water and molecular grade reagents
Tap water and deionized water contain a plethora of organic and inorganic impurities such as microorganisms, endotoxins, DNase and RNase, and salts. Your body easily protects you from these contaminants but they will wreck havoc on your molecular reactions. Thus, for all solutions that will come in contact with biological molecules use only 15 megohm (MΩ) water or distilled water.
For similar reasons, use only high quality molecular grade reagents (tested for the presence of DNases, RNases and Proteases).

Working with small volumes (1 µl to 1 ml).

  • When dispensing a small volume, touch the end of the pipette tip to the inside wall of the receiving tube to help draw the liquid out of the tip and into the tube.
  • Before opening tubes give the tube a 'quick spin'.
  • After setting up a 50 µl or smaller reaction, mix the reaction by pipetting up and down 2 or 3 times with a pipetter set to ~ 80% of the total volume (avoid foaming as many enzymes are destroyed by foaming). Then 'quick spin' the tube.

This technique pushes small volumes into a single drop
at the bottom of a tube where they are easier to find.
Place the tube and a balance tube (for volumes less than 10 µl you can use an empty tube) in the centrifuge and hold down the 'quick' button for 3 or 4 s. The drop should be at the bottom of the tube on the outside edge.

Using concentrated stock reagents

In molecular biology, we commonly work in relatively small volumes (less than a ml). This creates a problem for making solutions. For example, the working concentration for MgCl2 in Hind III digest is 10 mM.
The molecular weight of MgCl2 is 203.30, so to make 10 µl of 10 mM MgCl2 you would need

203.3 g/×L0.01M×0.000 01 L = 2.3×10-5 g

of MgCl2.

Instead of attempting to weigh out such small amounts, we make stock solutions at concentrations 10 times (10X) or greater than needed. We then simply add the appropriate volume to the reaction we are setting up, to get the desired final concentration.

For example, restriction enzyme buffers are usually made as 10X stocks. The working concentration is usually 1X. Therefore, you would dilute the stock buffer 1/10 into your reaction tube (e.g. if the final volume of the reaction is 10 µl, you would need to add 1 µl of the 10X stock buffer).

This means you need to regularly calculate dilution factors. For some practice problems see here.

Setting up several similar reactions – the easier way

When setting up several similar reactions use a master mix that contains all the common components, rather than setting up each reaction separately.
A master mix saves time and increases accuracy because you set up each reaction with only 2 pipetting steps, rather than 3 or more.

To set up a master mix, first work out how much of each component is added to a single digest, than multiply the amounts by the number of reactions + at least 10% for precision errors.

Table 1. Making a master mix. This master mix is for 5 reactions
where all components except the DNA are kept the same.

addition order component single reaction
volume to add
master mix
volume to add
DNA 5 0
2 10X Reaction buffer 1 5.5
3 BamH I (10 units/µl) 0.5 2.8
4 Hind III (10 units/µl) 0.5 2.8
1 H20 to 10 µl 3 16.5

Each reaction gets 5 µl of the master mix and 5 µl of the appropriate DNA.

Sterilizing in molecular biology

Most solutions we work with in molecular biology are sterile. The need for sterile growth media is obvious; however, most solutions used in molecular biology need to be kept sterile because they are excellent substrates for microbial growth. Even small amounts of microbial contamination can cause problems, especially if you will be using amplification techniques, such as PCR.

Keeping all solutions sterile means you can make larger batches of solutions to use over longer periods of time. Otherwise, you will find yourself preparing fresh solutions more often then you would like.

We also use only sterile consumables (e.g. tips and tubes) as biological contaminants can be carried on these.

Sterilization by autoclaving is an acceptable method for decontaminating biohazardous waste before disposal. We are required to decontaminate the following: all potentially hazardous organisms, genetically modified organisms, anything contaminated with such organisms, and nucleic acids from such organisms. In other words, pretty much everything we dispose of should be treated as biohazardous waste.

Autoclaving does not sufficiently decontaminate human or animal tissue; or volatile chemicals. Animal tissues must be incinerated. Volatile organics, such as phenol, or chloroform, must be sent to a certified chemical disposal plant.

Sterilizing by autoclaving

Autoclaving uses of pressurized steam to kill infectious agents and denature proteins. Water is heated to temperatures above its boiling point (to a minimum of 121 °C) by holding it in a pressurized chamber. This kind of "wet heat" is considered the most dependable method of sterilizing laboratory equipment and decontaminating biohazardous waste. Autoclaves do not remove chemical contamination. In fact autoclaving some chemicals can lead to production of dangerous fumes.

Packaging of items or liquids to be sterilized must permit heat (steam) penetration, and prevent pressure differentials that could result in breakage. This may be accomplished by using techniques such as:

  • loosening screw caps or using self venting caps,
  • capping open containers for sterilization with aluminum foil,
  • opening plastic bags prior to loading them into the autoclave.

Do not place sealed containers in an autoclave!
Additionally, make sure any containers you put in the autoclave can handle the heat and pressure. For glass, Pyrex is a good choice; for plastic use polypropylene (PP) or polycarbonate (PC).

Autoclaving liquids

Loosely set caps on bottles, or cover openings with foil or foam plugs.
Autoclave for 20 min at 120 °C and 15 lb/sq. in. using a slow exhaust (using fast exhaust can result in boiling over of superheated solutions). Longer times for autoclaving may be required if working with large volumes.
Only after solutions have completely cooled, tighten the lids.

Autoclaving plastics, tips in tip boxes, glass etc.

  • Wrap plastics, glass etc. in foil, PP bags, autoclavable plastic containers or Pyrex containers.
  • Loosely set caps on any containers.
  • Autoclave for 20 min at 120 °C and 15 lb/sq. in. Inert materials can be exhausted either fast or slow. Drying cycles are optional, if available they will remove most of the condensed water from the items.
  • Only after containers

have completely cooled, tighten the lids.

Filter sterilizing

Solutions that contain heat-labile components must be filter-sterilized. Such solutions include many proteins, vitamins, amino acids, carbohydrates and most antibiotics.

To filter sterilize small volumes (10 ml or less) use a filter which can be fitted on the end of a syringe. For larger volumes use sterile filters which can be fitted onto a bottle top. Porosity of the filter membrane should be no larger than 0.2 microns (µm). Collect the filtered solution in sterile containers.

Decontaminating solutions containing ethidium bromide

There are a variety of ways to remove ethidium bromide from solution. All of them involve binding the ethidium bromide with some type of solid. Once decontaminated, the remaining solution can be safely poured down the drain. The contaminated solid material is sent to a chemical waste disposal plant for further treatment.

Examples of solids that will bind ethidium bromide include:

  • activated charcoal
  • certain resins such as Amberlite XAD-16 (Sigma)

Decontaminating a liquid ethidium bromide solution with Amberlite XAD-16

(Lunn and Sansone 1987)

  1. Add 2.9 g of Amberlite XAD-16 for each 100 ml of solution up to 100 µg/ml ethidium bromide. Amberlite XAD-16, a nonionic, polymeric absorbent, is available from Rohm and Haas and sold by Sigma Chemical Company.
  2. Store the solution for 12 hours at room temperature, shaking it intermittently.
  3. Filter the solution through a Whatman No. 1 filter. Discard the filtrate to the drain.
  4. Seal the filter and Amberlite in a plastic bag and dispose of the bag through the Office of Research Safety.
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