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General Laboratory Methods


  • Safety Procedures
  • Preparation of Solutions
  • Disposal of Buffers and Chemicals
  • Equipment 
  • Micropipettors.
  • Using a pH meter
  • Autoclave operating procedures
  • Operating instructions for spectrophotometer
  • Working with DNA
  • Sterile Technique


    A number of chemicals used in the laboratory are hazardous. All manufacturers of hazardous materials are required by law to supply the user with pertinent information on any hazards associated with their chemicals. This information is supplied in the form of Material Safety Data Sheets or MSDS. This information contains the chemical name, CAS#, health hazard data, including first aid treatment, physical data, fire and explosion hazard data, reactivity data, spill or leak procedures, and any special precautions needed when handling this chemical. MSDS information can be accessed on World Wide Web. You are strongly urged to make use of this information prior to using a new chemical and certainly in the case of any accidental exposure or spill. The instructor or laboratory head must be notified immediately in the case of an accident involving any potentially hazardous reagents.

    The following chemicals are particularly noteworthy:

    These chemicals are not harmful if used properly: always wear gloves when using potentially hazardous chemicals and never mouth-pipet them. If you accidentally splash any of these chemicals on your skin, immediately rinse the area thoroughly with water and inform the instructor. Discard the waste in appropriate containers.


    Dosimeters must be worn when any radioactive work is performed. In addition, one is required to keep a record of wipe tests of radiation areas for every day that radioactivity is used. If any area registers greater than 200 dpm, that area must be immediately decontaminated and the area retested. The results of the original counting of the vials and any recounts should be placed in the appropriate section of a radiation log book with the date and the signature of the person conducting the test.

    A few common sense precautions should always be followed:


    Exposure to ultraviolet light can cause acute eye irritation. Since the retina cannot detect UV light, you can have serious eye damage and not realize it until 30 min to 24 hours after exposure. Therefore, always wear appropriate eye protection when using UV lamps.


    The voltages used for electrophoresis are sufficient to cause electrocution. Cover the buffer reservoirs during electrophoresis. Always turn off the power supply and unplug the leads before removing a gel.


    All common areas should be kept free of clutter and all dirty dishes, electrophoresis equipment, etc should be dealt with appropriately. Since you have only a limited amount of space to call your own, it is to your advantage to keep your own area clean. Since you will use common facilities, all solutions and everything stored in an incubator, refrigerator, etc. must be labeled. In order to limit confusion, each person should use his initials or other unique designation for labeling plates, etc. Unlabeled material found in the Gembox, incubators, or freezers may be destroyed. Always mark the backs of the plates with your initials, the date, and relevant experimental data, e.g. strain numbers. Each person will be assigned a general lab duty that may include keeping track of inventory, making sure a given area is kept clean, or maintaining equipment.



    1. A molar solution is one in which 1 liter of solution contains the number of grams equal to its molecular weight.
    Ex. To make up 100 ml of a 5M NaCl solution =
    58.456 (mw of NaCl) g x 5 moles x 0.1 liter = 29.29 g in water to a final volume of 100 ml
    2. Percent solutions.
    Percentage (w/v) = weight (g) in 100 ml of solution; Percentage (v/v) = volume (ml) in 100 ml of solution.
    Ex. To make a 0.7% solution of agarose in TBE buffer, weigh 0.7 g of agarose and bring up volume to 100 ml with TBE buffer.
    3. "X" Solutions.

    Many enzyme buffers are prepared as concentrated solutions, e.g. 5X or 10X (five or ten times the concentration of the working solution) and are then diluted such that the final concentration of the buffer in the reaction is 1X.

    Ex. To set up a restriction digestion in 25 ul, one would add 2.5 ul of a 10X buffer, the other reaction components, and water to a final volume of 25 ul.


    Many buffers in molecular biology require the same components but often in varying concentrations. To avoid having to make every buffer from scratch, it is useful to prepare several concentrated stock solutions and dilute as needed.

    Ex. To make 100 ml of TE buffer (10 mM Tris, 1 mM EDTA), combine 1 ml of a 1 M Tris solution and 0.2 ml of 0.5 M EDTA and 98.8 ml sterile water. The following is useful for calculating amounts of stock solution needed:

    Ci x Vi = Cf x Vf , where Ci = initial concentration, or conc of stock solution;
    Vi = initial vol, or amount of stock solution needed
    Cf = final concentration, or conc of desired solution;
    Vf = final vol, or volume of desired solution


    1. Refer to the laboratory manual for any specific instructions on preparation of the particular solution and the bottle label for any specific precautions in handling the chemical.

    2. Weigh out the desired amount of chemical(s). Use an analytical balance if the amount is less than 0.1 g.

    3. Place chemical(s) into appropriate size beaker with a stir bar.

    4. Add less than the required amount of water. Prepare all solutions with double distilled water.

    5. When the chemical is dissolved, transfer to a graduated cylinder and add the required amount of distilled water to achieve the final volume. An exception is in preparing solutions containing agar or agarose. Weigh the agar or agarose directly into the final vessel.

    6. If the solution needs to be at a specific pH, check the pH meter with fresh buffer solutions and follow instructions for using a pH meter.

    7. Autoclave, if possible, at 121oC for 20 min. Some solutions cannot be autoclaved, for example, SDS. These should be filter sterilized through a 0.22um filter. Media for bacterial cultures must be autoclaved the same day it is prepared, preferably within an hour or two. Store at room temperature and check for contamination prior to use by holding the bottle at eye level and gently swirling it.

    8. Solid media for bacterial plates can be prepared in advance, autoclaved, and stored in a bottle. When needed, the agar can be melted in a microwave, any additional components, e.g. antibiotics, can be added and the plates can then be poured.

    9. Concentrated solutions, e.g. 1M Tris-HCl pH=8.0, 5M NaCl, can be used to make working stocks by adding autoclaved double-distilled water in a sterile vessel to the appropriate amount of the concentrated solution.


    Glass and plastic ware used for molecular biology must be scrupulously clean. Dirty test tubes, bacterial contamination and traces of detergent can inhibit reactions or degrade nucleic acid.
    Glassware should be rinsed with distilled water and autoclaved or baked at 150oC for 1 hour. For experiments with RNA, glassware and solutions are treated with diethylpyrocarbonate to inhibit RNases which can be resistant to autoclaving.
    Plastic ware such as pipets and culture tubes are often supplied sterile. Tubes made of polypropylene are turbid and are resistant to many chemicals, like phenol and chloroform; polycarbonate or polystyrene tubes are clear and not resistant to many chemicals. Make sure that the tubes you are using are resistant to the chemicals used in your experiment. Micropipet tips and microfuge tubes should be autoclaved before use. Keep a supply of these for your own use.




        It is to everyone's advantage to keep the equipment in good working condition. As a rule of thumb, don't use anything unless you have been instructed in the proper use. This is true not only for equipment in the lab but also departmental equipment. Report any malfunction immediately. Rinse out all centrifuge rotors after use and in particular if anything spills. Please do not waste supplies - use only what you need. If the supply is running low, please notify either the instructor or the TA before the supply is completely exhausted. Occasionally, it is necessary to borrow a reagent or a piece of equipment from another lab. Except in an emergency, notify the instructor. Don't borrow anything on your own. In an emergency, please ask first.


        Most of the experiments you will conduct will depend on your ability to accurately measure volumes of solutions using micropipettors. The accuracy of your pipetting can only be as accurate as your pipettor and several steps should be taken to insure that your pipettes are accurate and are maintained in good working order.  They should then be checked for accuracy following the instructions given by the instructor. The simplest way is to use a calibrated pipet tip to see if the pipettor is dispensing the appropriate volume.  For Rainin pipetmen, the seals should be replaced every 6 months and the shaft cleaned regularly with isopropanol.  Since the pipettors will use different pipet tips, make sure that the pipet tip you are using is designed for your pipettor. DO NOT DROP IT ON THE FLOOR. If you suspect that something is wrong with your pipettor, first check the calibration to see if your suspicions were correct, then notify the instructor.


        Biological functions are very sensitive to changes in pH and hence, buffers are used to stabilize the pH. A pH meter is an instrument that measures the potential difference between a reference electrode and a glass electrode, often combined into one combination electrode. The reference electrode is often AgCl2. An accurate pH reading depends on standardization, the degree of static charge, and the temperature of the solution.

    1. The pH meter should be standardized each time it is used with a buffer of known pH, preferably one closest to the desired final pH. To calibrate the pH meter, expose the hole in the electrode, rinse the electrode with deionized water, and place the electrode in a standard solution, e.g., pH 7. Turn the selector to "pH". Adjust the pH meter to the appropriate pH. Turn selector to "standby". Rinse electrode with deionized water and place in a second standard buffer solution. The choice of the second standard depends on the final pH desired, for example, if the final pH desired is 8.5, the standard pH buffers used should be 7 and 10. If the final pH desired is 5.5, the standard pH buffers used should be 4 and 7. Turn the selector to "pH". Adjust the temperature knob to the second standard pH. Turn the selector to "standby" , rinse the electrode with deionized water, and return the electrode to the soaking solution. Cover up the hole in the electrode with the rubber gasket.

    2. When rinsing the electrode, never wipe the end with a Kimwipe but blot gently since wiping can create a static electric charge which can cause erroneous readings,

    3. Make sure the solution you are measuring is at room temperature since the pH can change with a change in temperature.

    4. Tris solutions cannot be measured with many pH meters. To prepare these solutions, use a Tris table that combines Tris base with Tris hydrochloride in varying proportions to achieve the desired pH.


    Place all material to be autoclaved in a autoclavable tray. All items should have indicator tape. Separate liquids from solids and autoclave separately. Make sure lids on all bottle are loose.
    1. Make sure chamber pressure is at 0 before opening the door.
    2. Place items to be autoclaved in the autoclave and close the door. Some autoclaves require that you also lock the door after it's closed.
    3. Set time - typically 20 minutes.
    4. Temperature should be set at 121oC already, but double-check and change if necessary.
    5. Set cycle: If liquid, set "liquid cycle" or "slow exhaust". If dry, set "dry cycle" or "fast exhaust" + dry time.
    6. Start the cycle. On some autoclaves, the cycle starts automatically at step 5. On others, turn to "sterilize".
    7. At the end of the cycle, check that a.) the chamber pressure is at 0 and b.) the temp is <100oC
    8. Open door. (On 3rd floor autoclave, don't push end cycle)
    9. Remove contents using gloves and immediately tighten all caps.


    To measure the absorbance of a solution in the short-wave range (<300 nM) use the quartz cuvettes. Disposable plastic cuvettes are available for reading in the visible range.

    Turn the spectrophotometer on - the switch is on the right in the back.

    Allow the instrument to calibrate. Do not open the chamber during this time. The deuterium lamp is OFF by default. To read absorbance in the UV range, turn the deuterium lamp on as follows after the machine has completed its calibration: Depress the function key until Fn5 is displayed. Press the mode key until d2on is displayed. Press enter. For best accuracy, the deuterium lamp should be warmed up for 20 minutes.

    Press the function key until Fn0 is displayed. Press enter. Using the up or down arrow keys, enter in the desired wavelength.

    Prepare a reference cuvette containing the same diluent as your sample. Prepare your sample.

    Place the reference cuvette in cell #1 and place your samples in cells #2-6.

    Press the cell key until cell #1 is in position. Press the Set Reference key to blank against the appropriate buffer. Press the cell key to advance to read the next sample.



        The following properties of reagents and conditions are important considerations in processing and storing DNA and RNA. Heavy metals promote phosphodiester breakage. EDTA is an excellent heavy metal chelator. Free radicals are formed from chemical breakdown and radiation and they cause phosphodiester breakage. UV light at 260 nm causes a variety of lesions, including thymine dimers and crosslinks. Biological activity is rapidly lost. 320 nm irradiation can also cause crosslinks, but less efficiently. Ethidium bromide causes photooxidation of DNA with visible light and molecular oxygen. Oxidation products can cause phosphodiester breakage. If no heavy metal are present, ethanol does not damage DNA. Nucleases are found on human skin; therefore, avoid direct or indirect contact between nucleic acids and fingers. Most DNases are not very stable; however, many RNases are very stable and can adsorb to glass or plastic and remain active. 5oC is one of the best and simplest conditions for storing DNA. -20oC: this temperature causes extensive single and double strand breaks. -70oC is probable excellent for long-term storage. For long-term storage of DNA, it is best to store in high salt (>1M) in the presence of high EDTA (>10mM) at pH 8.5. Storage of DNA in buoyant CsCl with ethidium bromide in the dark at 5oC is excellent. There is about one phosphodiester break per 200 kb of DNA per year. Storage of DNA in the phage is better than storing the pure DNA. [Davis, R.W., D. Botstein and J.R. Roth, A Manual for Genetic Engineering: Advanced Bacterial Genetics. Cold Spring Harbor Laboratories, Cold Spring Harbor, N.Y. 1980.]


    To remove protein from nucleic acid solutions:

    1. treat with proteolytic enzyme, e.g., pronase, proteinase K
    2. Phenol Extract. The simplest method for purifying DNA is to extract with phenol or phenol:chloroform and then chloroform. The phenol denatures proteins and the final extraction with chloroform removes traces of phenol. (E.3)
    3. CsCl/ethidium bromide density gradient


    1. Spectrophotometric. For pure solutions of DNA, the simplest method of quantitation is reading the absorbance at 260 nm where an OD of 1 in a 1 cm path length = 50 ug/ml for double-stranded DNA, 40 ug/ml for single-stranded DNA and RNA and 20-33 ug/ml for oligonucleotides. An absorbance ratio of 260 nm and 280 nm gives an estimate of the purity of the solution. Pure DNA and RNA solutions have OD260/OD280 values of 1.8 and 2.0, respectively. This method is not useful for small quantities of DNA or RNA (<1 ug/ml).
    2. Ethidium bromide fluorescence. The amount of DNA is a solution is proportional to the fluorescence emitted by ethidium bromide in that solution. Dilutions of an unknown DNA in the presence of 2 ug/ml ethidium bromide are compared to dilutions of a known amount of a standard DNA solutions spotted on an agarose gel or Saran Wrap or electrophoresed in an agarose gel.


    Precipitation with ethanol. (E.10) DNA and RNA solutions are concentrated with ethanol as follows: The volume of DNA is measured and the monovalent cation concentration is adjusted. The final concentration should be 2-2.5M for ammonium acetate, 0.3M for sodium acetate, 0.2M for sodium chloride and 0.8M for lithium chloride. The ion used often depends on the volume of DNA and on the subsequent manipulations; for example, sodium acetate inhibits Klenow, ammonium ions inhibit T4 polynucleotide kinase, and chloride ions inhibit RNA-dependent DNA polymerases. The addition of MgCl2 to a final concentration of 10mM assists in the precipitation of small DNA fragments and oligonucleotides. Following addition of the monovalent cations, 2-2.5 volumes of ethanol are added, mixed well, and stored on ice or at -20oC for 20 min to 1 hour. The DNA is recovered by centrifugation in a microfuge for 10 min (room temperature is okay). The supernatant is carefully decanted making certain that the DNA pellet, if visible, is not discarded (often the pellet is not visible until it is dry). To remove salts, the pellet is washed with 0.5-1.0 ml of 70% ethanol, spun again, the supernatant decanted, and the pellet dried. Ammonium acetate is very soluble in ethanol and is effectively removed by a 70% wash. Sodium acetate and sodium chloride are less effectively removed. For fast drying, the pellet can spun briefly in a Speedvac, although the method is not recommended for many DNA preparations as DNA that has been overdried is difficult to resuspend and also tends to denature small fragments of DNA. Isopropanol is also used to precipitate DNA but it tends to coprecipitate salts and is harder to evaporate since it is less volatile. However, less isopropanol is required than ethanol to precipitate DNA and it is sometimes used when volumes must be kept to a minimum, e.g., in large scale plasmid preps.


    Restriction and DNA modifying enzymes are stored at -20oC in a non-frost free freezer, typically in 50% glycerol. The tubes should never be allowed to reach room temperature and gloves should be worn when handling as fingers contain nucleases. Always use a new, sterile pipet tip every time you use a restriction enzyme. Also, the volume of the enzyme should be less than 1/10 of the final volume of the reaction mixture.


    1. All media, including plates, liquid media and top agar must be autoclaved immediately after it is prepared. It is best to prepare media in several small bottles, only opening one at a time. Check the bottle for contamination before you use it by gently swirling it and looking for cloudy material in the center. Always grow up a small amount of broth alone when growing cells overnight. A small amount of contamination is not always evident until the media is incubated at 37oC.

    2. Use a flame on inoculating loops and on the lips of media bottles before and after pipetting from them. Never leave a media or agar bottle open on the bench and don's take an individually-wrapped pipet out of its protective wrapper until you are ready to use it (i.e., don't walk across the room with an unwrapped pipet). Always use a fresh, sterile pipet or pipet tip when pipetting culture media, and never go back into a media bottle or cell culture with a used pipet.

    3. To prevent wide-scale, untraceable contamination, each person should have his own stock of liquid culture media, top agar, plates, 100% glycerol, glycerol stocks of cells, etc. and don't share.

    4. Overnight cultures should be grown only from a single colony on a fresh plate or from a previously-tested glycerol stock that was grown from a single colony. To prepare an overnight culture from a glycerol stock, take an individually-wrapped 1-ml pipet and a culture tube of media to the -80oC freezer. Quickly remove the cap from the freezer vial containing the glycerol stock, scrap a small amount of ice from the surface of the culture, replace the cap on the freezer vial, and place the pipet into the culture tube. Sufficient numbers of bacteria are present in the ice in order for the culture to grow to saturation in 16 hours. Never let the glycerol stock thaw.

    4. Think about what you are doing. The best defense is common sense.