Molecular Biology Laboratory Manual

Molecular Biology
Laboratory Manua

By Denny R Randall

General Laboratory Procedures, Equipment Use, and Safety
Considerations

I. Safety Procedures

A. Chemicals

A number of chemicals used in any molecular biology 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. A file containing MSDS information on the hazardous substances should
be kept in the lab. In addition, 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/lab manager must be notified immediately in the case of an accident
involving any potentially hazardous reagents.

The following chemicals are particularly noteworthy:

• Phenol - can cause severe burns

• Acrylamide - potential neurotoxin

• Ethidium bromide - carcinogen

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.

B. Ultraviolet Light

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.

C. Electricity

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.

D. General Housekeeping

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 refrigerators, 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.

General Laboratory Procedures, Equipment Use, and Safety
Considerations

II. Preparation of Solutions

A. Calculation of Molar, % and "X" Solutions .

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 NaCI solution
= 58.456 (mw of NaCI) g/mol x 5 moles/liter x 0.1 liter = 29.29 g in 100 ml of
solution

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, weight 0.7 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
IX. Ex. To set up a restriction digestion in 25 u I, one would add 2.5 u I of a 10X
buffer, the other reaction components, and water to a final volume of 25 u I.

B. Preparation of Working Solutions from Concentrated Stock Solutions .

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: CixVi = CfxVf, where
C i = initial concentration, or cone of stock solution; V i = initial vol, or amount of
stock solution needed C f = final concentration, or cone of desired solution; V f =
final vol, or volume of desired solution

C. Steps in Solution Preparation:

1. Refer to a laboratory reference manual for any specific instructions on
preparation of the particular solution and the bottle label for any specific
precautions in handling the chemical. Weigh out the desired amount of
chemical(s). Use an analytical balance if the amount is less than 0.1 g. Place
chemical(s) into appropriate size beaker with a stir bar. Add less than the
required amount of water. Prepare all solutions with double distilled water

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. 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. Autoclave, if possible, at 121 deg C for 20
min. Some solutions cannot be autoclaved, for example, SDS. These should
be filter sterilized through a 0.22 u m or 0.45 u m 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 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.
2. Concentrated solutions, e.g. 1M Tris-HCI pH=8.0, 5M NaCI, 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.

D. Glassware and Plastic Ware .

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 150
degrees C for 1 hour. For experiments with RNA, glassware and solutions are
treated with diethyl-pyrocarbonate 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. Micro pipet tips and
microfuge tubes should be autoclaved before use.

General Laboratory Procedures, Equipment Use, and Safety
Considerations

III. Disposal of Buffers and Chemicals

1. Any uncontaminated, solidified agar or agarose should be discarded in the
trash, not in the sink, and the bottles rinsed well.

2. Any media that becomes contaminated should be promptly autoclaved
before discarding it. Petri dishes and other biological waste should be
discarded in Biohazard containers which will be autoclaved prior to disposal.

3. Organic reagents, e.g. phenol, should be used in a fume hood and all organic
waste should be disposed of in a labeled container, not in the trash or the
sink.

4. Ethidium bromide is a mutagenic substance that should be treated before
disposal and should be handled only with gloves. Ethidium bromide should
be disposed of in a labeled container.

5. Dirty glassware should be rinsed, all traces of agar or other substance that
will not come clean in a dishwasher should be removed, all labels should be
removed (if possible), and the glassware should be placed in the dirty dish
bin. Bottle caps, stir bars and spatulas should not be placed in the bins but
should be washed with hot soapy water, rinsed well with hot water, and
rinsed three times with distilled water.

General Laboratory Procedures, Equipment Use, and Safety
Considerations

IV. Equipment

A. General Comments

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/lab managerbefore 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.

B. Micropipettors

Most of the experiments you will conduct in this laboratory 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. Each pair of students will be assigned a set of pipettors and
upon receipt, they should be labeled with the students' name. They should then be
checked for accuracy following the instructions given by the instructor. If they
need to be recalibrated, do so. We have two different types of pipettors, Rainin
pipetmen and Oxford benchmates. 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.

C. Using a pH Meter

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 AgCI 2. An
accurate pH reading depends on standardization, the degree of static charge, and
the temperature of the solution.

Operation of Orion PerpHecT pH Meter

• Expose hole on side of electrode by sliding the collar down. Make sure there
is sufficient electrode filling solution in the electrode (it should be up to the
hole). If not, fill with ROSS filling solution only (Do not use any filling
solution containing silver (Ag).

• Ensure that sample to be pHed is at room temperature and is stirring gently
on the stir plate.

• Calibrate the pH meter with the two solutions that bracket the target pH - 4
and 7 or 7 and 10 as follows:

• Press the CAL key to initialize the calibration sequence. The last calibration
range will be displayed (e.g. 7-4). Press YES to accept or use the scroll keys
to select a different range. Press YES to accept.

• The number 7 will light up on the left hand side of the screen indicating that
the meter is ready to accept the pH 7 standard buffer. Rinse off electrode
and place in fresh pH 7 standard buffer solution. The READY light will come
on when the value has stabilized. Press YES to accept the value.

• The number 4 (or 10) will light up next indicating that the meter is ready to
accept the pH 4 (or 10) standard buffer solution. Rinse off electrode and
place in fresh pH 4 standard buffer solution. The READY light will come on
when the value has stabilized. Press YES to accept the value.

• SLP will be displayed. The meter will then go MEASURE mode.

• Rinse electrode and place into sample. The READY light is displayed when
signal is stable.

D. Autoclave Operating Procedures

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. Do not crowd large number of items in tray- in order for
all items to reach the appropriate temperature, one must allow sufficient
air/steam circulation.

1. Make sure chamber pressure is at 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 121 deg C 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; b. the
temp is <100 deg C

8. Open door.

9. Remove contents using gloves and immediately tighten all caps.

E. Operating Instructions for Spectrophotometer - Pharmacia Ultraspec

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 FnO 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.

. PROPER USE OF THE MICROSCOPE

1 . When moving your microscope, always carry it with both hands (Figure 1 ). Grasp
the arm with one hand and place the other hand under the base for support.

2. Turn the revolving nosepiece so that the lowest power objective lens is "clicked"
into position.

3. Place the microscope slide on the stage and fasten it with the stage clips. You can
push down on the back end of the stage clip to open it.

4. Using the coarse adjustment, lower the objective lens down as far as it will go
without touching the slide! Note: Look at the slide and lens from the side when doing
this.

5. Look through the eyepiece and adjust the illuminator (or mirror) and diaphragm for
the greatest amount of light.

6. Slowly turn the coarse adjustment so that the objective lens goes up (away from the
slide). Continue until the image comes into focus. Use the fine adjustment, if available,
for fine focusing.

7. Move the microscope slide around so that the image is in the center of the field of
view and readjust the mirror, illuminator or diaphragm for the clearest image.

8. You should be able to change to the next objective lenses with only slight focusing
adjustment. Use the fine adjustment, if available. If you cannot focus on your
specimen, repeat steps 4 through 7 with the higher power objective lens in place. DO
NOT ALLOW THE LENS TO TOUCH THE SLIDE!

9. The proper way to use a monocular microscope is to look through the eyepiece with
one eye and keep the other eye open (this helps avoid eye strain). If you have to close
one eye when looking into the microscope, it's ok. Remember, everything is upside
down and backwards. When you move the slide to the right, the image goes to the
left!

10. Do not touch the glass part of the lenses with your fingers. Use only special lens
paper to clean the lenses.

1 1 . When finished, raise the tube, click the low power lens into position and remove
the slide

General Laboratory Procedures, Equipment Use, and Safety
Considerations

V. Working with DNA

A. Storage

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
cross-link. Biological activity is rapidly lost. 320 nm irradiation can also cause
cross-link, but less efficiently. Ethidium bromide causes photo oxidation 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. 5 E C is one of the best and simplest conditions for storing DNA. -20
deg C: this temperature causes extensive single and double strand breaks. -70 E C
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 CsCI with ethidium bromide in the dark at 5 E C is
excellent. There is about one phosphodiester break per 200 kb of DNA per year.
Storage of A DNA in the phage is better than storing the pure DNA. [ ref : 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.]

B. Purification

To remove protein from nucleic acid solutions:

1. Treat with proteolytic enzyme, e.g., pronase, proteinase K

2. Purify on a silica-based column such as a Qiagen PCR Prep Column

3. CsCI/ethidium bromide density gradient

4. Phenol Extract. The simplest method for purifying DNA is to extract with
phenol or phenokchloroform and then chloroform. The phenol denatures
proteins and the final extraction with chloroform removes traces of phenol

5. Purify on silica-based column such as Qiagen Brand columns
(http://www.qiagen.com)

C. Quantitation .

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 u g/ml for double-stranded DNA, 40 u g/ml for single-
stranded DNA and RNA and 20-33 u g/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 OD 260/OD 280 values of
1.8 and 2.0, respectively. This method is not useful for small quantities of
DNA or RNA (<1 u g/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 u g/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.

D. Concentration

Precipitation with ethanol. 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 MgCI 2 to a final concentration of lOmM 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 -20 E C 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 over dried 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.

E. Restriction Enzymes

Restriction and DNA modifying enzymes are stored at -20 deg C in a non-frost free
freezer, typically in 50% glycerol. The enzymes are stored in an insulated cooler
which will keep the enzymes at -20 deg C for some period of time. 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

General Laboratory Procedures, Equipment Use, and Safety
Considerations

VI. Sterile Technique

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 37 deg C.

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 -80 deg
C 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.

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

General Laboratory Procedures, Equipment Use, and Safety
Considerations

VII. Working with E. coli

A. Small Scale Cultures

Experiments using E. coli cells should always be done on fresh cultures, either
from a freshly streaked plate or from a glycerol stock. To grow a small scale E. coli
culture, prepare 3-5 ml of LB (or appropriate broth - include antibiotic if the
culture contains a plasmid) in two sterile 50 ml tubes. (Note: smaller tubes can be
used but the culture will not be appropriately aerated and hence will not grow well
and is not recommended). Inoculate one tube with a single colony from a fresh
plate or a scraping from a glycerol stock. The second tube is used as a broth
control. Incubate both tubes at 37 deg C, shaking vigorously overnight. Inspect
the tubes the next morning. The broth control should be clear and the inoculated
culture should be very turbid. Make a note of any debris found in the tubes and
only incubate longer if the culture is not dense. Do not allow cells to overgrow.
Use immediately. For some applications, cells can be stored at 4 deg C for short
periods prior to use.

B. Permanent Storage

For every culture used, in particular, for newly constructed strains or for cells
containing plasmids, a permanent glycerol stock must be prepared as soon as the
construct has been confirmed and this stock must be placed in the laboratory
stock collection with the appropriate documentation and location information.
Failure to follow these procedures will result in serious penalties. This procedure
pertains not only to E. coli but to any organism for which a deep freeze stock can
be prepared. Also, all plasmid constructs, including construction intermediates,
must be maintained in cells, not as naked DNA stocks. For each construct, at least
2 stocks should be made. To prepare a glycerol stock for E. coli cells, combine 1.4
ml of a freshly grown overnight culture with 0.6 ml of sterile 50% glycerol. Mix
well. Transfer to two freezer vials labeled with the strain name, the date and your
initials (not an eppendorf tube). Immediately place into a dry ice/ethanol bath or
into a box in the -80 deg C freezer. Note the location and enter data into the strain
book.

Instructions for Notebook Keeping

1. A notebook should be kept for laboratory experiments only using a Scientific
Notebook Co. book or other bound book I personally like the cheap collage ruled
books you can find at wall mart are the dollar store. The notebook should be
written in ink, and each page signed and dated. Mistakes are not to be erased but
should be marked out with a single line. Try to keep your notebook with the idea
that someone else must be able to read and understand what you have done. The
notebook should always be up-to-date and can be collected at any time. I also
keep scan each page and save them to a DVD as well as a cheap sd card. Your sd
cards DVDs and notebooks should cost no more then 20 to 30 dollars' per
experiment set.

2. INDEX: An index containing the title of each experiment and the page number
should be included at the beginning of the notebook.

3. WHAT SHOULD BE INCLUDED IN THE NOTEBOOK? Essentially

everything you do in the laboratory should be in your notebook as well as
breaks and bathroom breaks. The notebook should be organized by experiment
only and should not be organized as a daily log. Start each new experiment on a
new page. The top of the page should contain the title of the experiment, the date,
and the page number. The page number is important for indexing, referring to
previous experiments, and for labeling materials used in a given experiment. If an
experiment spans more than one page, note the page on which the experiment
continues if it's not on the next page. Each experiment should include the
following:

4. Title/Purpose: Every experiment should have a title and it should be
descriptive. An example would be"

Method and techniques for investigating a phenomenon in
The honey bee know as Colony collapse disorder"

When starting a new project, it is a good idea to introduce the overall strategy
prior to beginning the first experiment. This serves two purposes. First, it forces
you to think about what you are doing and why and sometimes things look
differently when written down than they do in your head. Second, ideas can be
patented, and a thorough description of your hypothesis and experimental
strategy with appropriate documentation can be helpful for any future intellectual
property issues.

5. Background information: This section should include any information that
is pertinent to the execution of the experiment or to the interpretation of the
results. For example, if it is a repeat experiment, state what will be done
differently to get the experiment to work. If it's a cloning experiment, include
what the strategy is and how the recombinants will be screened. A simple drawing
of the plasmid map can be helpful. This is not like the introduction to a paper.
Include anything that will be helpful in carrying out the experiment and
deciphering the experiment at a later date. For the most part, notebooks are not
written for today but for the future.

6. Materials: This section should include the key materials, i.e., solutions or
equipment, that will be needed. It is not necessary to include every piece of lab
equipment required, i.e. vortexer, pipetman, etc, but you should include any
specialized equipment and the manufacturer, i.e, a phosphoimager or real-time
PCR instrument. Composition of all buffers should be included unless they are
standard or are referenced. Pre-packaged kits should be identified as to the name
of the kit, the vendor, and the catalog number. Biological samples should be
identified by genus and species, strain number, tissue type, and/or genotype with
the source of the material identified. Enzymes should be identified by name,
vendor, and concentration. DNA samples should be identified as to 1: type of DNA,
i.e., chromosomal, plasmid, etc, 2: purity (miniprep, gel purified, PCR product) 3:
concentration, if known, and 4: source, (include prior experiment number if the
DNA was isolated in a previous experiment). Include all calculations made in

preparing solutions. The sequence of all oligonucleotides must be included or
referenced. Agarose gels should be identified by percentage and buffer used. If
any of these materials were used in previous experiments, include only the
reference to that earlier experiment, do not repeat the information again.

7. Procedure: Write down exactly what you are going to do before you do it and
make sure you understand each step before you do it. In general, Xerox copies of
procedures are not acceptable for several reasons: 1. You should include
everything you do including all volumes and amounts; many protocols are written
for general use and must be adapted for a specific application. 2. Writing a
procedure out helps you to remember and to understand what it is about. It will
also help you to identify steps that may be unclear or that need special attention.
3. Some procedures can be several pages long and include more information than
is necessary in a notebook. However, it is good laboratory practice to have a
separate notebook containing methods that you use on a regular basis (this is not
required for this course). If an experiment is a repeat of an earlier experiment,
you do not have to write down each step but refer to the earlier experiment by
page or experiment number. If you make any changes, note the changes and why.
Flow charts are sometimes helpful for experiments that have many parts. Tables
are also useful if an experiment includes a set of reactions with multiple variables.
It is good practice to check off steps as they are completed or reagents as they are
added to prevent you from losing you place or for forgetting to add something. All
procedures should be referenced.

8. Results: This section should include all raw data, including gel photographs,
printouts, colony counts, autoradiographs, etc. All lanes on gel photographs must
be labeled and always identify the source and the amount of any standards. This
section should also include your analyzed data, for example, transformation
efficiencies, calculations of specific activities or enzyme activities.

9. Conclusions/Summary: This is one of the most important sections. You
should summarize all of your results, even if they were stated elsewhere and state
any conclusions you can make. If the experiment didn't work, what went wrong
and what will you do the next time to try to trouble shoot?

Common Stock Solutions

10 M Ammonium Acetate

To prepare a 10 M solution in 100 ml, dissolve 77 g of ammonium acetate in 70 ml of H 2 at
room temperature. To prepare a 5 M solution in 100 ml, dissolve 38.5 g in 70 ml of H 2 0.
Adjust the volume to 100 ml with H 2 0. Sterilize the solution by passing it through a 0.22pm
filter. Store the solution in tightly sealed bottles at 4 ° C or at room temperature.
Ammonium acetate decomposes in hot H 2 and solutions containing it should not be
autoclaved.

Ampicillin

Prepare a stock of 100 mg/ml in water. Sterilize by filtration. Store at -20°C but avoid
repeated freeze/thaw cycles. Use at a final concentration of 100 ug/ml.

Cresol Red Loading Dye - 2.5X - for PCR reactions

1M sucrose, 0.02% cresol. Prepare 1% cresol red in water (0.5 g/50 ml). To prepare loading
dye, dissolve 17 g sucrose in a total volume of 49 ml of water. Add 1 ml 1% cresol red.

EB buffer (Qiagen recipe)

10 mMTris-CI pH 8.3

EDTA stock

To prepare 1 liter, 0.5M EDTA pH 8.0: Add 186.1 g of disodium EDTA-2H 20 to 800 ml of H
20. Stir vigorously on a magnetic stirrer. Adjust the pH to 8.0 with NaOH (approx. 20 g of
NaOH pellets). Dispense into aliquots and sterilize by autoclaving. The disodium salt of
EDTA will not go into solution until the pH of the solution is adjusted to approx. 8.0 by the
addition of NaOH. For tetrasodium EDTA, use 226.1 g of EDTA and adjust pH with HCI.

6x gel loading buffer

0.25% Bromophenol blue
0.25%Xylene cyanol FF
15% Ficoll Type 4000
120 mM EDTA

IPTG

IPTG is isopropylthio-b-D-galactoside. Make a 20% (w/v, 0.8 M) solution of IPTG by
dissolving 2 g of IPTG in 8 ml of distilled H 20. Adjust the volume of the solution to 10 ml
with H 20 and sterilize by passing it through a 0.22pm disposable filter. Dispense the
solution into 1 ml aliquots and store them at -20 ° C.

LB Medium

To make 1 liter, use 10 g tryptone, 5 g yeast extract, 10 g NaCI. Adjust pH to 7.0. Sterilize
by autoclaving.

LB Agar

Dispense 15 g per liter of agar directly into final vessel. Prepare LB medium as above and
add to agar. NOTE: Agar will not go into solution until it is autoclaved (or boiled). If adding
antibiotics, autoclave medium first and allow to cool until warm to the touch, then add the
antibiotic. Dispense about 30 ml per plate. Allow plates to dry either at 37°C overnight or
20 minutes in a laminar flow hood (lids removed). Store in original Petri plate bags,
inverted, at 4°C for up to 2 weeks.

NaCI

To prepare 1 liter of a 5 M solution: Dissolve 292 g of NaCI in 800 ml of H 20. Adjust the
volume to 1 liter with H 20. Dispense into aliquots and sterilize by autoclaving. Store the
NaCI solution at room temperature.

NaOH

The preparation of 10 N NaOH involves a highly exothermic reaction, which can cause
breakage of glass containers. Prepare this solution with extreme care in plastic beakers. To
800 ml of H20, slowly add 400g of NaOH pellets, stirring continuously. As an added
precaution, place the beaker on ice. When the pellets have dissolved completely, adjust the
volume to 1 liter with H20. Store the solution in a plastic container at room temperature.
Sterilization is not necessary.

20X SB (electrophoresis buffer)

(Buffer diluted to IX should be 10 mM Sodium hydroxide and pH 8.5 )

for 1 liter, weigh out 8 g NaOH and ~40 g boric acid - add water, dissolve and add
additional boric acid until pH = 8.0; bring final volume to 1 liter.

SDS stock

10% or 20% (w/v) SDS. Also called sodium lauryl (or dodecyl) sulfate. To prepare a 20%
(w/v) solution, dissolve 200 g of electrophoresis-grade SDS in 900 ml of H 2 0. Heat to 68 °
C and stir with a magnetic stirrer to assist dissolution. If necessary, adjust the pH to 7.2 by

adding a few drops of concentrated HCI. Adjust the volume to 1 liter with H 20. Store at
room temperature. Sterilization is not necessary. Do not autoclave.

Use a mask when weighing this out.

20x SSC

0.3M Na(3) citrate
3M NaCI

SOB Medium: per liter:

Bacto-tryptone 20 g

Yeast extract 5 g

NaCI 0.584 g

KCI 0.186 g Mix components and adjust pH to 7.0 with NaOH and autoclave.

2 M Mg ++ stock:

MgCI 2-6H20 20.33 g
MgSO 4 -7H20 24.65 g

Distilled water to 100 ml. Autoclave or filter sterilize.

2 M Glucose

Glucose 36.04 g

Distilled water to 100 ml. Filter sterilize.

For SOB Medium + magnesium: Add 1 ml of 2 M Mg ++ stock to 99 ml SOB
Medium.

For SOCMedium: Add 1 ml of 2 M Mg ++ stock and 1 ml of 2 M Glucose to 98 ml of
SOB Medium.

3M Sodium Acetate - pH 5.2

To prepare a 3 M solution: Dissolve 408.3 g of sodium acetate-3H 2 in 800 ml of H 20.
Adjust the pH to 5.2 with glacial acetic acid. Adjust the volume to 1 liter with H 2 0. Dispense
into aliquots and sterilize by autoclaving.

Southern Solutions:

Depurination solution (for Southern blotting)

250mM HCI
Denaturation solution (for Southern blotting)

1.5M NaCI
0.5M NaOH

Neutralization solution (for Southern blotting)

1.5M NaCI

0.5M Tris-HCI, pH adjusted to 7.5

50x TAE

Prepare a 50x stock solution in 1 liter of H 2 0:
242 g of Tris base
57.1 ml of glacial acetic acid
100 ml of 0.5 M EDTA (pH 8.0)

The lx working solution is 40 mM Tris-acetate/1 mM EDTA.

5X (or 10X) TBE

Prepare a 5x stock solution in 1 liter of H20:
54 g of Tris base
27.5 g of boric acid
20 ml of 0.5 M EDTA (pH 8.0)

The pH of the concentrated stock buffer should be approx. 8.3.. Some investigators prefer
to use more concentrated stock solutions of TBE (lOx as opposed to 5x). However, 5x stock
solution is more stable because the solutes do not precipitate during storage. Passing the 5x
or lOx buffer stocks through a 0.22pm filter can prevent or delay formation of precipitates.

TE buffer:

10 mM Tris-CI (pH, usually 7.6 or 8.0)
1 mM EDTA (pH 8.0)

Use concentrated stock solutions to prepare. If sterile water and sterile stocks are used,
there is no need to autoclave. Otherwise, sterilize solutions by autoclaving for 20 minutes.
Store the buffer at room temperature.

1 M Tris-CI - used at various pHs

Using Tris base : To make 1 liter, dissolve 121 g Tris Base in 800 ml of water. Adjust pH
to the desired value by adding approximately the following:

pH = 7.4 about 70 ml of concentrated HCI

pH = 7.6 about 60 ml of concentrated HCI
pH = 8.0 about 42 ml of concentrated HCI

Make sure solution is at room temperature before making final pH adjustments.
Bring final volume to 1 liter. Sterilize by autoclaving.

Using Trizma tables: an alternate procedure for preparing Tris solutions is to
combine the proper amount of Tris Base and Tris Hydrochloride to achieve the
desired value using Sigma's Tris tables.

WB (10% redistilled glycerol, 90% distilled water, v/v)

In a 1-liter graduated cylinder, add 100 ml of glycerol and 900 ml of distilled water.
Cover with parafilm and mix thoroughly. Sterilized by autoclaving, and chill to 4°C.

Western Blotting Solutions:

IX Transfer buffer 1: 25 mM Tris, 192 mM Glycine, pH 8.3

Mix 3.03 g Tris, 14.4 g glycine; add dd water to 1 liter - do not adjust pH.

IX Transfer buffer 2: 25 mM Tris, 192 mM Glycine, pH 8.3, 20 % methanol

Mix 3.03 g Tris, 14.4 g glycine; add 200 ml methanol; add dd water to 1 liter - do
not adjust pH. (NOTE: methanol is not needed for PVDF membranes)

10X Western Buffer: 200 mM Tris pH = 7.5; 1.5 M NaCI

To prepare IX Western Buffer, dilute 10X buffer to IX, adding Tween-20 to 0.1%.
Remove 50 ml and set aside for the last two washes. To the remainder, add I-Block
to 0.2%, heating gently with constant stirring until dissolved. Bring to room
temperature before using.

X-gal 5-bromo-4-chloro-3-indolyl-b-D-galactoside (same recipe for X-phosphate)

Make a 2% (w/v) stock solution by dissolving X-gal in dimethylformamide at a
concentration of 20 mg/ml solution. Use a glass or polypropylene tube. Wrap the
tube containing the solution in aluminum foil to prevent damage by light and store at
-20 ° C. It is not necessary to sterilize X-gal solutions.

• Preparation of Genomic DNA from Bacteria- using Phase
Lock Gel™

(Modified from Experimental Techniques in Bacterial Genetics, Jones and Bartlet, 1990)

• Materials: see Solutions for Recipes

IE buffer

10% (w/v) sodium dodecyl sulfate (SDS)
•20 mg/ml proteinase K
phenol\chloroform (50:50)
isopropanol
70% ethanol

3M sodium acetate pH 5.2
Phase Lock Gel™ (Eppendorf-Brinkmann)

PCR Amplification of DNA

Materials:

sterile water

10X amplification buffer with 15mM MgCI2

10 mM dNTP

50 pM oligonucleotide primer 1

50 uM oligonucleotide primer 2

5 unit/pl Taq Polymerase

template DNA (1 ug genomic DNA, 0.1-1 ng plasmid DNA) in 10 pi

mineral oil (for thermocyclers without a heated lid

1. Combine the following for each reaction (on ice) in a 0.2 or 0.5 ml
tube:

10X PCR buffer

10 pi

Primer 1

1 Ml

Primer 2

1 Ml

dNTP

2 pi

template DNA and water

85.5 pi

Taq Polymerase

0.5 pi

2. Prepare a control reaction with no template DNA and an additional 10
pi of sterile water.

3. If the thermocycler does not have a heated lid, add 70-100 pi mineral
oil (or 2 drops of silicone oil) to each reaction.

4. Place tubes in a thermal cycler preheated to 94 degrees C.

5. Run the following program:

94 degrees C 1 min

55 degrees C 1 min or annealing temperature appropriate for
particular primer pair

72 degrees C 1 min (if product is <500 bp), 3 min (if product is
>500 bp)
for 30 cycles.
Program a final extension at 72 degrees C for 7 min

Restriction Enzyme Digestion of DNA

Materials:

•lOX restriction enzyme buffer (see manufacturer's

recommendation)
•DNA

sterile water

restriction enzyme

phenokchloroform (1:1) (optional)

• Add the following to a microfuge tube:

•2 ul of appropriate 10X restriction enzyme buffer

•0.1 to 5 ug DNA
sterile water to a final volume of 19 ul (Note: These volumes are
for analytical digests only. Larger volumes may be necessary for
preparative digests or for chromosomal DNA digests.

• Add 1 to 2 ul (3 to 20 units) enzyme and mix gently. Spin for a
few seconds in microfuge.

• Incubate at the appropriate temperature (usually 37 degrees
C) for 1 to 2 hours.

. Run a small aliquot on a gel to check for digestion.

• If the DNA is to be used for another manipulation, heat inactivate the
enzyme (if it is heat labile) at 70 degrees C for 15 min,
phenol/chloroform extract and ethanol precipitate, or purify on Qiagen
DNA purification column

Phenol/chloroform Extraction of DNA

Materials:

phenokchloroform (1:1)
chloroform

• Add an equal volume of buffer-saturated phenokchloroform (1:1) to
the DNA solution.

• Mix well. Most DNA solutions can be vortexed for 10 sec except for
high molecular weight DNA which should be gently rocked. (If using
Phase-Lock Gel, follow procedure MJJ

• Spin in a microfuge for 3 min.

• Carefully remove the aqueous layer to a new tube, being careful to
avoid the interface. (Steps 1-4 can be repeated until an interface is no
longer visible).

• To remove traces of phenol, add an equal volume of chloroform to the
aqueous layer.

• Spin in a microfuge for 3 min.

• Remove aqueous layer to new tube.

• Ethanol precipitate the DNA

Ethanol Precipitation of DNA

Materials: see Solutions for Recipes

3 M sodium acetate pH 5.2 or 5 M ammonium acetate
•DNA

• 100% ethanol

• Measure the volume of the DNA sample. Adjust the salt concentration by adding
1/10 volume of sodium acetate, pH 5.2, (final concentration of 0.3 M) or an equal
volume of 5 M ammonium acetate (final concentration of 2.0-2.5 M). These amounts
assume that the DNA is in TE only; if DNA is in a solution containing salt, adjust salt
accordingly to achieve the correct final concentration. Mix well. Add 2 to 2.5 volumes
of cold 100% ethanol (calculated after salt addition). Mix well.

• Place on ice or at -20 degrees C for ^20 minutes.

• Spin a maximum speed in a microfuge 10-15 min. Carefully decant supernatant. Add
1 ml 70% ethanol. Mix. Spin briefly. Carefully decant supernatant. Air dry or briefly
vacuum dry pellet.

• Resuspend pellet in the appropriate volume of TE or water

Agarose Gel Electrophoresis

Materials:

agarose solution in TBE , TAE or SB (generally 0.7-1%)

• IX TBE, TAE, or SB (same buffer as in agarose)
gel loading dye

•10 mg/ml ethidium bromide

• To prepare 100 ml of a 0.7% agarose solution, measure 0.7 g agarose into a glass
beaker or flask and add 100 ml IX buffer. Microwave or stir on a hot plate until
agarose is dissolved and solution is clear.

• Allow solution to cool to about 55 degrees C before pouring. (Ethidium bromide can
be added at this point to a concentration of 0.5 ug/ml)

• Prepare gel tray by sealing ends with tape or other custom-made dam. Place comb in
gel tray about 1 inch from one end of the tray and position the comb vertically such
that the teeth are about 1-2 mm above the surface of the tray

• Pour 50 degree C gel solution into tray to a depth of about 5 mm. Allow gel to
solidify about 20 minutes at room temperature.

• To run, gently remove the comb, place tray in electrophoresis chamber, and cover
(just until wells are submerged) with electrophoresis buffer (the same buffer used to
prepare the agarose). Excess agarose can be stored at room temperature and
remelted in a microwave. To prepare samples for electrophoresis, add 1 ul of 6x gel
loading dye for every 5 pi of DNA solution. Mix well. Load 5-12 pi of DNA per well

(for minigel). Electrophorese at 50-150 volts until dye markers have migrated an
appropriate distance, depending on the size of DNA to be visualized.
• If the gel was not stained with ethidium during the run, stain the gel in 0.5 ug/ml
ethidium bromide until the DNA has taken up the dye and is visible under short wave
UV light, if the DNA will not be used further, or with a hand-held long-wave UV light
if the DNA is to be cut out and purified.

Transformation of E. coli by Electroporation

(electroporation procedure from Cell-Porator™ Voltage Booster, Life
Technologies, Cat. Series 1612)

Background:

There are two methods to transform E. coli cells with plasmid DNA -
chemical transformation and electroporation. For chemical
transformation, cells are grown to mid-log phase, harvested and
treated with divalent cations such as CaCI 2 . Cells treated in such a
way are said to be competent. To chemically transform cells,
competent cells are mixed with the DNA , on ice, followed by a brief
heat shock. Then, cells are incubated with rich medium and allowed
to express the antibiotic resistant gene for 30-60 minutes prior to
plating. For electroporation, cells are also grown to mid-log phase
but are then washed extensively with water to eliminate all salts.
Usually, glycerol is added to the water to a final concentration of
10% so that the cells can be stored frozen and saved for future
experiments. To electroporate DNA into cells, washed E. coli are
mixed with the DNA to be transformed and then pipetted into a
plastic cuvette containing electrodes. A short electric pulse, about
2400 volts/cm, is applied to the cells causing smalls holes in the
membrane through which the DNA enters. The cells are then
incubated with broth as above before plating.

For chemical transformation, there is no need to pre-treat the DNA.
For electroporation, the DNA must be free of all salts so the
ligations are first precipitated with alcohol before they are used.

Experimental Design :

To determine the efficiency of transformation, a positive control
transformation should be done using 1 ng of uncut plasmid DNA,
e.g. pUC19. The efficiency of transformation is calculated as the

number of transformants/ug of input DNA. A negative control should
also be included that contains cells with no added DNA.

A negative control with cells only (no added DNA) should also be
included.

Host Cells

For most cloning applications, we use DH5a host cells. These cells
are compatible with lacZ blue/white selection procedures, are easily
transformed, and good quality plasmid DNA can be recovered from
transformants. One notable exception is when transforming with
plasmid constructs containing recombinant genes under control of
the T7 polymerase. These constructs are typically transformed into
DH5a for the cloning phase, but need to be transformed into a
different bacterial strain, BL21(DE3) for expression of the
recombinant protein (BL21 strains carry the gene for expression of
the T7 polymerase).

Electroporation of E. coli:

Materials:

■ Sterile centrifuge bottles - 250 ml for GSA rotor

■ SOB medium

■ E. coli host strain such as DH5a

■ WB (10% redistilled glycerol, 90% distilled water, v/v) chilled to 4°C-
need 500 ml of WB for each 250 ml of culture

■ tRNA (5-10 pg/ml - used as a mass carrier to increase the efficiency of
precipitation)

■ 5 M ammonium acetate

■ 100% ethanol

■ 70% ethanol

■ 0.5XIE or EB (10 mM Tris, pH 8.3)

■ SOC medium

■ transformation plates

I. Preparation of E. coli cells for electroporation.

1. Use a fresh colony of DH5a (or other appropriate host strain) to
inoculate 5 ml of SOB (without magnesium) medium in a 50 ml

sterile conical tube. Grow cells with vigorous aeration overnight at
37°C.

2. Dilute 2.5 ml of cells into 250 ml of SOB (without magnesium) in
a 1 liter flask. Grow for 2 to 3 hours with vigorous aeration at 37°C
until the cells reach an OD 550 = 0.8.

3. Harvest cells by centrifugation at 5000 RPM in a GSA rotor for 10
min in sterile centrifuge bottles. (Make sure you use autoclaved
bottles!).

4. Wash the cell pellet in 250 ml of ice-cold WB as follows. First, add
a small amount of WB to cell pellet; pipet up and down or gently
vortex until cells are resuspended. Then fill centrifuge bottle with ice
cold WB and gently mix. NOTE- the absolute volume of WB added at
this point is not important.

5. Centrifuge the cell suspension at 5,000 RPM for 15 min and
carefully pour off the supernatant as soon as the rotor stops. Cells
washed in WB do not pellet well. If the supernatant is turbid,
increase the centrifugation time.

6. Wash the cell pellet a second time by resuspending in 250 ml of
sterile ice-cold WB using the same technique described above.
Centrifuge the cell suspension at 5000 RPM for 15 min.

7. Gently pour off the supernatant leaving a small amount of WB in the
bottom of the bottle. Resuspend the cell pellet in the WB - no additional WB
needs to be added - and the final volume should be about 1 ml. Cells can be
used immediately or can be frozen in 0.2 ml aliquots in freezer vials using a
dry ice-ethanol bath. Store frozen cells at -70°C.

II. Preparing DNA for Electroporation

DNA for electroporation must have a very low ionic strength and a
high resistance. The DNA may be purified by either dilution,
precipitation or dialysis.

• For transformation of purified plasmid DNA, dilute DNA in 10
mM Tris pH 8-8.3 to about 1-50 ng/ul (do not use TE). Use 1
ul for transformation.

. For ligation reactions, use the following procedure.
Purifying DNA by Precipitation:

1. Add 5 to 10 ug of tRNA to a 20 ul ligation reaction in a 1.5 ml
tube. Add 22 ul 5M ammonium acetate (or an equal volume of
ligation reaction with added tRNA). Mix well.

2. Add 100 ul absolute ethanol (or 2.5 volumes of ligation reaction,
tRNA and salt). Ice 15 min.

3. Centrifuge at >12,000 x g for 15 min at 4°C. Carefully decant the
supernatant.

4. Wash the pellet with 1 ml of 70% ethanol. Centrifuge at > 12,000
x g for 15 min at room temperature. Remove the supernate.

5. Air dry the pellet (speed vac okay but don't overdry).

6. Resuspend the DNA in EB buffer (10 mM Tris-HCI, pH 8.3) or
0.5X TE buffer [5 mM Tris-HCI, 0.5 mM EDTA (pH 7.5)] to a
concentration of 10 ng/ul of DNA. For ligation reactions, it is
convenient to resuspend in 10 ul. Use 1 ul per transformation of 20
ul of cell suspension.

III. Electroporation.

1. Mark the required number of micro centrifuge tubes. Place the
required number of Micro-electroporation Chambers on ice. Fill the
temperature control compartment of the Chamber Safe with ~250
ml of ice-water slurry and place the Chamber Rack in the Chamber
Safe.

2. Thaw an aliquot of cells that have prepared as in Section I and
aliquot 20 ul of cells to the required number of microfuge tubes on
ice. Add 1 ul of the DNA (or ligation reaction) prepared as in Section
II.

3. Using a micro pipette, pipette 20 ul of the cell-DNA mixture
between the bosses in a Micro-Electroporation Chamber. Do not
leave an air bubble in the droplet of cells; the pressure of a bubble

may cause arcing and loss of the sample. Place the chamber in a
slot in the Chamber Rack and note its position. Repeat the process if
more than one sample is to be pulsed. Up to 4 samples can be
placed in the Chamber Rack at one time. Handle the chambers
gently to avoid accidentally displacing the sample from between the
bosses.

4. Close the lid of the Chamber safe and secure it with the draw
latch.

5. Plug the pulse cable into the right side of the Chamber safe.

6. Turn the chamber selection knob on top of the Chamber Safe to
direct the electrical pulse to the desired Micro-Electroporation
Chamber.

7. Set the resistance on the Voltage Booster to 4 kQ; set the Pulse
Control unit to LOW and 330 uF; double check connections.

8. Charge the Pulse Control unit by setting the CHARGE ARM switch
on the Pulse Control unit to CHARGE and then pressing the UP
voltage control button until the voltage reading is 5 to 10 volts
higher than the desired discharge voltage. For E. co\\, the standard
conditions are 2.4 kv, which means setting the Pulse Control unit to
405 volts (400 volts is the desired discharge voltage + 5). The
voltage booster amplifies the volts by ~6-fold such that the total
discharge voltage is 2400 volts, or 2.4 kv. The actual peak voltage
delivered to the sample will be shown on the Voltage Booster meter
after the pulse is delivered.

9. Set the CHARGE/ARM switch to the ARM position. The green light
indicates that the unit is ready to deliver a DC pulse. Depress the
pulse discharge TRIGGER button and hold for 1 second.

NOTE: The DC voltage display on the Pulse Control unit should read
<10 volts after a pulse has been delivered. If not, discharge the
capacitor using the DOWN button.

10. For additional samples, turn the chamber selection knob to the
next desired position and repeat steps 8 and 9 until all samples are
pulsed.

. 11. For ampicillin selection, inoculate the samples into 2 ml of
SOC medium and shake for 30-60 minutes to allow expression
of the antibiotic gene. Plate cells on LB medium with
appropriate antibiotic or screening reagent (e.g. 100 ug/ml
ampicillin, and/or 40 ul of 20 mg/ml X-Gal, XP, and 40 ul of
100 mM IPTG).

Preparative DNA Fragment Isolation from an Agarose Gel

Background:

DNA can be easily isolated and purified after size selection on an agarose
gel. The fragment of interest is simply cut out of the gel with a razor blade
and purified by a number of different methods.The easiest is to use a
method that involves first dissolving the agarose slice in a solution at 50°C,
then binding the DNA from the melted agarose to a silica-gel membrane.

1. Prepare an agarose gel in TAE buffer using the four-well combs.
(Preparative agarose gels should be run using IX TAE electrophoresis
and gel buffer as the borate in TBE interferes with some purification
resin). Load the DNA. To visualize the DNA after staining, do not
expose the DNA to shortwave UV light as this will introduce
nicks. Visualize the bands with a hand-held long wave UV light and
cut out the band with a clean razor blade (Note: place gel on a glass
slide to avoid cutting the surface of the transilluminator).

2. After cutting out the band, follow the procedure for DNA fragment
purification using Qiagen QIAquick or Qiaex II purification systems
following the manufacturer's procedure (" http://www.qiaqen.com") .
Estimate the approximate concentration of the DNA obtained by
running 10% of the eluate on an agarose gel against a DNA mass
ladder.

Ligations of plasmid DNA to insert DNA

A typical ligation reaction consists of about 20-200 ng of a vector
and a 1-3 fold molar excess of insert DNA. A typical ligation reaction
consists of the following:

. 20-100 ng vector

. 3-fold MOLAR excess (not mass) of insert DNA

. 4 |_j I 5X ligase buffer or 2 ul 10X ligase buffer (note: ligase
buffer contains ATP and is unstable when repeatedly frozen
and thawed. Prepare small aliquots of buffer and discard
aliquot after use)

. water to 19 ul

. 1 [i\ T4 DNA ligase

Incubate at room temperature for 2-24 hours. For transformation by
electroporation, ethanol precipitate as described in the
transformation procedure.

Procedure for Transfection of Mammalian Cells

Materials:

Lipofectamine (Invitrogen)

IMDM containing 10% fetal bovine serum, 1% glutamine, 1% aa
•IMDM containing 1% glutamine
IMDM containing 20% fetal bovine serum, 1% glutamine, 1% aa

1. In a six-well or 35 mm tissue culture plate, seed ~2x 10 5 cells per well in 2 ml IMDM
containing 10% FBS and nonessential amino acids.

2. Incubate the cells at 37°C in a C0 2 incubator until the cells are 70-80% confluent.
This will usually take 18-24 h.

3. Prepare the following solutions in 12 x 75 mm sterile tubes:

Solution A: For each transfection, dilute 2 ug DNA (plasmid) in 375 pi serum-free
IMDM (containing nonessential amino acids).

Solution B: For each transfection, dilute 12 pi LIPOFECTAMINE Reagent in 375 pi
serum-free IMDM.

4. Combine the two solutions, mix gently, and incubate at room temperature for 15-45
min. The solution may appear cloudy, however this will not impede the
transfection. Wash the cells once with 2 ml serum-free IMDM.

5. For each transfection, add 750 pi serum-free IMDM to each tube containing the lipid-
DNA complexes. Do not add antibacterial agents to media during transfection. Mix
gently and overlay the diluted complex solution onto the washed cells.

6. Incubate the cells for 5 h at 37°C in a C0 2 incubator.

7. Add 1.5 ml IMDM with 20% FBS without removing the transfection mixture. If
toxicity is a problem, remove the transfection mixture and replace with normal
growth medium. Replace medium at 18-24 h following start of transfection.

8. Assay cell extracts for gene activity 24-72 h after the start of transfection, depending
on cell type and promoter activity.

Southern Blotting

1. Electrophoresis of DNA is carried out in a neutral agarose gel system. Prepare a 0.8-
1% agarose gel containing lx TAE buffer. Ethidium bromide can be added to a final
concentration of 0.2 ug/ml.

2. Apply the samples to the gel.

3. Run the gel in lx TAE. buffer at 4V/cm until the bromophenol blue indicates that the
sample has run for a sufficient distance.

4. Following electrophoresis, visualize the gel under UV transillumination and
photograph with a ruler.

5. i) Depurination, 10 minutes at room temperature with gentle agitation (optional).
This step is necessary if target sequences are greater than 10 Kb in size

ii) Denaturation, 25 minutes at room temperature with gentle agitation,
iii) Neutralization, 30 minutes at room temperature with gentle agitation. When using
nitrocellulose membranes, the neutralization time should be extended to 45 minutes.
Include a rinse in distilled water between each step

6. Assemble the capillary blotting apparatus using 10X SSC as the transfer buffer. Allow
the DNA to transfer overnight onto Hybond N + .

7. The following day, disassemble the apparatus, mark the membrane appropriately
and fix the DNA to the membrane by UV crosslinking or baking (2 hours at 80°C).
For nitrocellulose membranes, bake for 2 hrs. at 80°C in a vacuum oven.

Solutions :
Hybridization buffer

5x SSC

1 in 20 dilution Liquid Block (Amersham) or other blocking reagent

0.1% (w/v) SDS

5%(w/v) Dextran sulphate
EDTA stock

0.5M EDTA pH8.0
SDS stock

10% or 20% (w/v) SDS
Depurination solution

250mM HCI
Denaturation solution

1.5M NaCI

0.5M NaOH
Neutralization solution

1.5M NaCI

0.5M Tris-HCI

pH adjusted to 7.5
20x SSC

0.3M Na(3) citrate

3M NaCI

Western Blot Analysis of Epitoped-tagged Proteins Using The
Chemifluorescent Detection Method - for alkaline
phosphatase conjugated antibodies

1. Cut PVDF membrane to the appropriate size, activate with absolute methanol for 5
sec, and incubate in distilled water for 5 min.

2. For electroblotting, equilibrate in transfer buffer and follow the standard blotting
procedure to transfer the proteins to the membrane. For dot blotting, keep
membrane wet until ready to use.

3. After protein has been transferred to the membrane, wash again in absolute
methanol for a few seconds and allow to dry at room temperature for 30 min. or
more.

4. Block in 30 ml of IX Western buffer (containing 0.1% Tween-20 and 0.2% I-Block),
gently rocking, 1 hr, room temperature.

5. Add appropriate dilution of primary antibody (typically 1:5000 or 1:10,000) prepared
in IX Western buffer (containing 0.1% Tween-20 and 0.2% I-Block), incubate 30
min, room temperature, gently rocking.

6. Wash three times in 20 ml IX Western buffer (containing 0.1% Tween-20 and 0.2%
I-Block) for 5 min each. Add appropriate dilution of secondary antibody conjugated
to alkaline phosphatase prepared in IX Western buffer (containing 0.1% Tween-20
and 0.2% I-Block), gently rocking, 30 min, room temperature.

7. Wash as in step #6.

8. Then, wash twice with IX Western buffer without I-block.

9. At the end of the second final wash, leave some buffer in the container to keep the
membrane moist. With the membrane facing protein-side up, add 0.5 ml of substrate
solution directly into the remaining liquid, mix well, and pipet (with a plOOO) the
solution over the membrane to ensure the entire surface comes into contact with the
substrate. Gently agitate for a few minutes, remove membrane to a paper towel and
let dry completely. The substrate solution can be reused immediately for additional
membranes.

10. Scan membrane using the Molecular Dynamics Storm or other suitable instrument.

Western Blotting Solutions:

IX Transfer buffer: 25 mM Tris, 192 mM Glycine, pH 8.3. Mix 3.03 g Tris and 14.4 g
glycine; add water to 1 liter - do not add acid or base to pH - it should be >8.0. Use 0.5X
for transfer in 20% methanol.

10X Western Buffer: 200 mM Tris pH = 7.5; 1.5 M NaCI (containing 0.1% Tween-20
and 0.2% I-Block). To prepare IX Western Buffer, dilute 10X buffer to IX, adding Tween-
20 to 0.1%. Remove 50 ml and set aside for the last two washes. To the remainder, add
I-Block to 0.2% (Cat #T2015, Applied Biosystems - formerly Tropix). To dissolve I-Block,
heat solution in a beaker briefly in a microwave to about 60°C, then stir until dissolved
(solution will be cloudy). Bring to room temperature before using.
Primary antibody: For his tagged proteins - Anti-His monoclonal antibody - BD
Bioscience #631212

'Secondary antibody: Goat anti-mouse alkaline phosphatase conjugated - Biorad #170-
6520.

Substrate: ECF chemifluorescent substrate - Amersham #RPN5785. Mix substrate with
accompanying buffer as per manufacturer's recommended instructions, prepare 1 ml
aliquots and store at -20°C.

Recommended Cycle Sequencing Protocols For ABI 3100
Template Quantity

Template

Quantity

PCR product:

100-200bp

200-500bp

500-lOOObp

1000-2000bp

>2000bp

1-3 ng
3-10 ng
5-20 ng
10-40 ng
20-50 ng

Single-stranded

25-50 ng

Double-stranded

150-300 ng

Cosmid, BAC

0.5-1.0 mg

Bacterial genomic DNA

2-3 mg

Reaction Mixtures

Reagent

Full Reaction

Half Reaction

Half Reaction(10 m>)

Big Dye Premix

8 Ml

4 Ml

2 Ml

Big Dye Seq.
Buffer

_

2 Ml

1 Ml

Template

See Table above

See Table above

See Table above

Primer (10 pM)

1 Ml

1 Ml

1 Ml

Water

q.s.

q.s.

q.s.

Total

20 Ml

20 Ml

10 Ml

Primer Quantity

Primer needed = 3.2 - 10 pmoles

PCR Cycle Sequencing Settings for Big Dye V3.1

Initial denaturing

96°C lmin

25 cycles of

96°C lOsec

50°C 5sec

60°C 4min

Hold at

4°C

Ethanol/EDTA Precipitation to clean up reactions

• Add 5uL of 125 mM EDTA. Make sure the EDTA reaches the bottom of
the tube.

Add 60uL of 100% ethanol to each tube.

Finger vortex and incubate at room temperature for 15 min.

Spin samples in a microcentrifuge at max speed in 4 ° C for 20min.

Carefully aspirate off the supernatant.

Add 60 pi of 70% Ethanol.

Spin samples in a microcentrifuge at maximum speed at 4 ° C for 15

minutes.

Aspirate off the supernatant.

Dry sample for 15 minutes in a Speed-Vac (longer if air-drying).

Protect samples from light while they are drying.

Colony PCR

1. Prepare a master mix containing the following:
IX reaction:

. 2.5 pi 2 mM dNTPs

• 2.5 pi 10 pM primer 1

• 10 pi 2.5X cresol red loading dye
. 2.5 pi 10X PCR buffer

• 5.0 pi sterile water

• 0.5-1 pi Taq polymerase

Note: If using PuReTaq Ready-To-Go PCR Beads (Amersham
Biosciences), use 10 pi of water instead of dNTP's, buffer and enzyme]

To prepare the master mix, multiply the volumes above by the number
of colonies to be screened + 1.

One-Step Gene Assembly

2. Aliquot to 0.2 ml labeled PCR tubes and keep on ice.

3. Prepare one selection plate, e.g., LB + ampicillin, for every 20 colonies
screened.

4. Label the plate with a grid so that each colony can be associated with
a number that matches the number on the PCR tube and can be
retrieved once PCR results are known.

5. With a toothpick or sterile loop, pick colonies from transformation
plate, patch onto the selection plate and then place remainder in PCR
with the same identifier.

PCR cycle using the same conditions for the original PCR with those
same primers with one modification: include a 5 min 94°C
denaturation at the beginning of the cycling reaction.
Analyze PCR results by running reactions directly onto an agarose gel
(no additional loading dye is required)

5.
6.

Design overlapping oligonucleotides, generally 40 bases in length, that
encompass the sense strand of the gene of interest. Design antisense
oligonucleotides that stagger the sense oligos by 20 bases.
Design outside PCR amplification primers to incorporate the
appropriate restriction enzyme recognition sites, if desired, and to
overlap the assembled gene sequence by at least 15 nucleotides.
(Order oligonucleotides from IDT or Invitrogen using their standard
desalting purification).

Reconstitute oligonucleotides to 100 uM in 10 mM Tris pH=8.5 (same
as Qiagen buffer EB). Vortex well to reconstitute and store at -20°C.
Prepare a gene assembly mix by combining 5 pi of each of the gene
assembly oligos. Dilute this mix so that each oligo is at a final
concentration of 1 pM. (This will be referred to as IX). Then dilute this
mix 1:2 (0.5X), 1:5 (0.2X) and 1:10 (0.1X) in Tris buffer. This step is
to optimize the assembly/amplification of the required product which
varies from one gene to the next - so best to set up 4 different
reactions, one for each dilution of the mix, so determine which gives
you the best yield and the least background.
Prepare 10 pM dilutions of outside amplification primers.
Perform one step gene assembly/amplifications using the following:

Reaction conditions:

Template

5 pi gene assembly mix (IX, 0.5X, 0.2X, and
0.1X)

Primers (0.4 uM final)

2 pi of 10 pM outside amplification primer #1
(OP-5')

2 pi of 10 pM outside amplification primer #2
(OP-3')

dNTP (0.2 mM final)

5 pi of 2 mM each (provided with KOD
enzymes)

10X PCR buffer

5 pi (provided with KOD enzymes)

25 mM MgCI2

2 pi for KOD HiFi; none needed for XL

Sterile water

28 pi for KOD HiFi; 30 pi for XL

KOD HiFi enzyme for
highest accuracy or

0.4 pi KOD HiFi (Novagen) or

KOD XLforTA cloning

1 pi for KOD XL (Novagen)

Total

50 Ml

Cycling conditions:

KOD XL- up to 2 kb

KOD HiFi - up to 2 kb

25 cycles

94°C

30 sec

98°C

15 sec

52°C

5 sec

52°C

2 sec

72°C

30-60 sec/kb

72°C

20 sec

1 cycle

74°C

10 min

• Analyze 5ul on agarose gel.

• Purify remainder using Qiagen PCR purification columns, digest
overnight, then gel purify. Ligate to vector of choice.

Tissue Culture Methods

I. TYPES OF CELLS GROWN IN CULTURE

Tissue culture is often a generic term that refers to both organ culture and cell culture
and the terms are often used interchangeably. Cell cultures are derived from either
primary tissue explants or cell suspensions. Primary cell cultures typically will have a
finite life span in culture whereas continuous cell lines are, by definition, abnormal and
are often transformed cell lines.

II. WORK AREA AND EQUIPMENT

A. Laminar flow hoods. There are two types of laminar flow hoods, vertical and
horizontal. The vertical hood, also known as a biology safety cabinet, is best for working
with hazardous organisms since the aerosols that are generated in the hood are filtered
out before they are released into the surrounding environment. Horizontal hoods are
designed such that the air flows directly at the operator hence they are not useful for
working with hazardous organisms but are the best protection for your cultures. Both
types of hoods have continuous displacement of air that passes through a HEPA (high
efficiency particle) filter that removes particulates from the air. In a vertical hood, the
filtered air blows down from the top of the cabinet; in a horizontal hood, the filtered air
blows out at the operator in a horizontal fashion. NOTE: these are not fume hoods and
should not be used for volatile or explosive chemicals. They should also never be used
for bacterial or fungal work. The hoods are equipped with a short-wave UV light that can
be turned on for a few minutes to sterilize the surfaces of the hood, but be aware that
only exposed surfaces will be accessible to the UV light. Do not put your hands or face
near the hood when the UV light is on as the short wave light can cause skin and eye
damage. The hoods should be turned on about 10-20 minutes before being used. Wipe
down all surfaces with ethanol before and after each use. Keep the hood as free of
clutter as possible because this will interfere with the laminar flow air pattern.

B. C0 2 Incubators. The cells are grown in an atmosphere of 5-10% C0 2 because the

medium used is buffered with sodium bicarbonate/carbonic acid and the pH must be
strictly maintained. Culture flasks should have loosened caps to allow for sufficient gas
exchange. Cells should be left out of the incubator for as little time as possible and the
incubator doors should not be opened for very long. The humidity must also be
maintained for those cells growing in tissue culture dishes so a pan of water is kept filled
at all times.

C. Microscopes. Inverted phase contrast microscopes are used for visualizing the cells.
Microscopes should be kept covered and the lights turned down when not in use. Before
using the microscope or whenever an objective is changed, check that the phase rings
are aligned.

D. Preservation. Cells are stored in liquid nitrogen (see Section III- Preservation and
storage).

E. Vessels. Anchorage dependent cells require a nontoxic, biologically inert, and
optically transparent surface that will allow cells to attach and allow movement for
growth. The most convenient vessels are specially-treated polystyrene plastic that are
supplied sterile and are disposable. These include petri dishes, multi-well plates,
microtiter plates, roller bottles, and screwcap flasks - T-25, T-75, T-150 (cm 2 of surface
area). Suspension cells are either shaken, stirred, or grown in vessels identical to those
used for anchorage-dependent cells.

III. PRESERVATION AND STORAGE. Liquid N 2 is used to preserve tissue culture cells,
either in the liquid phase (-196°C) or in the vapor phase (-156°C). Freezing can be
lethal to cells due to the effects of damage by ice crystals, alterations in the
concentration of electrolytes, dehydration, and changes in pH. To minimize the effects of
freezing, several precautions are taken. First, a cryoprotective agent which lowers the
freezing point, such as glycerol or DMSO, is added. A typical freezing medium is 90%
serum, 10% DMSO. In addition, it is best to use healthy cells that are growing in log
phase and to replace the medium 24 hours before freezing. Also, the cells are slowly
cooled from room temperature to -80°C to allow the water to move out of the cells
before it freezes. The optimal rate of cooling is 1°-3°C per minute. Some labs have
fancy freezing chambers to regulate the freezing at the optimal rate by periodically
pulsing in liquid nitrogen. We use a low tech device called a Mr. Frosty (C#1562 -
Nalgene, available from Sigma). The Mr. Frosty is filled with 200 ml of isopropanol at
room temperature and the freezing vials containing the cells are placed in the container
and the container is placed in the -80°C freezer. The effect of the isopropanol is to allow
the tubes to come to the temperature of the freezer slowly, at about 1°C per minute.
Once the container has reached -80°C (about 4 hours or, more conveniently, overnight)
the vials are removed from the Mr. Frosty and immediately placed in the liquid nitrogen
storage tank. Cells are stored at liquid nitrogen temperatures because the growth of ice
crystals is retarded below -130°C. To maximize recovery of the cells when thawing, the
cells are warmed very quickly by placing the tube directly from the liquid nitrogen
container into a 37°C water bath with moderate shaking. As soon as the last ice crystal
is melted, the cells are immediately diluted into prewarmed medium.

IV. MAINTENANCE

Cultures should be examined daily, observing the morphology, the color of the medium
and the density of the cells. A tissue culture log should be maintained that is separate

from your regular laboratory notebook. The log should contain: the name of the cell line,
the medium components and any alterations to the standard medium, the dates on
which the cells were split and/or fed, a calculation of the doubling time of the culture
(this should be done at least once during the semester), and any observations relative
to the morphology, etc.

A. Growth pattern. Cells will initially go through a quiescent or lag phase that depends
on the cell type, the seeding density, the media components, and previous handling.
The cells will then go into exponential growth where they have the highest metabolic
activity. The cells will then enter into stationary phase where the number of cells is
constant, this is characteristic of a confluent population (where all growth surfaces are
covered).

B. Harvesting. Cells are harvested when the cells have reached a population density
which suppresses growth. Ideally, cells are harvested when they are in a semi-confluent
state and are still in log phase. Cells that are not passaged and are allowed to grow to a
confluent state can sometime lag for a long period of time and some may never recover.
It is also essential to keep your cells as happy as possible to maximize the efficiency of
transformation. Most cells are passaged (or at least fed) three times a week.

1. Suspension culture. Suspension cultures are fed by dilution into fresh medium.

2. Adherent cultures. Adherent cultures that do not need to be divided can simply be fed
by removing the old medium and replacing it with fresh medium.

When the cells become semi-confluent, several methods are used to remove the cells
from the growing surface so that they can be diluted:

• Mechanical - A rubber spatula can be used to physically remove the cells from
the growth surface. This method is quick and easy but is also disruptive to the
cells and may result in significant cell death. This method is best when harvesting
many different samples of cells for preparing extracts, i.e., when viability is not
important.

• Proteolytic enzymes - Trypsin, collagenase, or pronase, usually in combination
with EDTA, causes cells to detach from the growth surface. This method is fast
and reliable but can damage the cell surface by digesting exposed cell surface
proteins. The proteolysis reaction can be quickly terminated by the addition of
complete medium containing serum

• EDTA - EDTA alone can also be used to detach cells and seems to be gentler on
the cells than trypsin. The standard procedure for detaching adherent cells is as
follows:

1. Visually inspect daily

2. Release cells from monolayer surface

a. wash once with a buffer solution

b. treat with dissociating agent

c. observe cells under the microscope. Incubate until cells become
rounded and loosen when flask is gently tapped with the side of the hand.

d. Transfer cells to a culture tube and dilute with medium containing

serum.

e. Spin down cells, remove supernatant and replace with fresh medium.

f. Count the cells in a hemacytometer, and dilute as appropriate into fresh
medium.

C. Media and growth requirements

1. Physiological parameters

A. temperature - 37C for cells from homeother

B. pH - 7.2-7.5 and osmolality of medium must be maintained

C. humidity is required

D. gas phase - bicarbonate cone, and C0 2 tension in equilibrium

E. visible light - can have an adverse effect on cells; light induced production of
toxic compounds can occur in some media; cells should be cultured in the dark
and exposed to room light as little as possible;

2. Medium requirements: (often empirical)

A. Bulk ions - Na, K, Ca, Mg, CI, P, Bicarb or C0 2

B. Trace elements - iron, zinc, selenium

C. sugars - glucose is the most common

D. amino acids - 13 essential

E. vitamins - B, etc.

F. choline, inositol

G. serum - contains a large number of growth promoting activities such as buffering
toxic nutrients by binding them, neutralizes trypsin and other proteases, has undefined
effects on the interaction between cells and substrate, and contains peptide hormones
or hormone-like growth factors that promote healthy growth.

H. antibiotics - although not required for cell growth, antibiotics are often used to
control the growth of bacterial and fungal contaminants.

3. Feeding - 2-3 times/week.

4. Measurement of growth and viability. The viability of cells can be observed visually
using an inverted phase contrast microscope. Live cells are phase bright; suspension
cells are typically rounded and somewhat symmetrical; adherent cells will form
projections when they attach to the growth surface. Viability can also be assessed using
the vital dye, trypan blue, which is excluded by live cells but accumulates in dead cells.
Cell numbers are determined using a hemocytometer.

V. SAFETY CONSIDERATIONS

Assume all cultures are hazardous since they may harbor latent viruses or other
organisms that are uncharacterized. The following safety precautions should also be
observed:

pipetting: use pipette aids to prevent ingestion and keep aerosols down to a minimum
•no eating, drinking, or smoking
wash hands after handling cultures and before leaving the lab
decontaminate work surfaces with disinfectant (before and after)
autoclave all waste

use biological safety cabinet (laminar flow hood) when working with hazardous
organisms. The cabinet protects worker by preventing airborne cells and viruses
released during experimental activity from escaping the cabinet; there is an air barrier
at the front opening and exhaust air is filtered with a HEPA filter make sure cabinet is
not overloaded and leave exhaust grills in the front and the back clear (helps to
maintain a uniform airflow)
use aseptic technique
dispose of all liquid waste after each experiment and treat with bleach

REFERENCES:

R. Ian Freshney, Culture of Animal cells: A manual of basic techniques, Wiley-Liss,
1987.

VI. TISSUE CULTURE PROCEDURES

Each student should maintain his own cells throughout the course of the experiment.
These cells should be monitored daily for morphology and growth characteristics, fed
every 2 to 3 days, and subcultured when necessary. A minimum of two 25 cm 2 flasks
should be carried for each cell line; these cells should be expanded as necessary for the
transfection experiments. Each time the cells are subcultured, a viable cell count should
be done, the subculture dilutions should be noted, and, after several passages, a
doubling time determined. As soon as you have enough cells, several vials should be
frozen away and stored in liquid N 2 . One vial from each freeze down should be thawed
1-2 weeks after freezing to check for viability. These frozen stocks will prove to be vital
if any of your cultures become contaminated.

Procedures: 1. Media preparation. Each student will be responsible for maintaining his
own stock of cell culture media; the particular type of media, the sera type and
concentration, and other supplements will depend on the cell line. Do not share media
with you partner (or anyone else) because if a culture or a bottle of media gets
contaminated, you have no back-up. Most of the media components will be purchased
prepared and sterile. In general, all you need to do is sterily combine several sterile
solutions. To test for sterility after adding aN components, pipet several mis from each
media bottle into a small sterile petri dish or culture tube and incubate at 37EC for
several days. Use only media that has been sterility tested. For this reason, you must
anticipate your culture needs in advance so you can prepare the reagents necessary.
But, please try not to waste media. Anticipate your needs but don't make more than you
need. Tissue culture reagents are very expensive; for example, bovine fetal calf serum
cost ~ $200/500 ml. Some cell culture additives will be provided in a powdered form.
These should be reconstituted to the appropriate concentration with double-distilled
water (or medium, as appropriate) and filtered (in a sterile hood) through a 0-22 um
filter.

All media preparation and other cell culture work must be performed in a laminar flow
hood. Before beginning your work, turn on blower for several minutes, wipe down all

surfaces with 70% ethanol, and ethanol wash your clean hands. Use only sterile pipets,
disposable test tubes and autoclaved pipet tips for cell culture. All culture vessels, test
tubes, pipet tip boxes, stocks of sterile eppendorfs, etc. should be opened only in the
laminar flow hood. If something is opened elsewhere in the lab by accident, you can
probably assume its contaminated. If something does become contaminated,
immediately discard the contaminated materials into the biohazard container and notify
the instructor.

2. Growth and morphology. Visually inspect cells frequently. Cell culture is sometimes
more an art than a science. Get to know what makes your cells happy. Frequent feeding
is important for maintaining the pH balance of the medium and for eliminating waste
products. Cells do not typically like to be too confluent so they should be subcultured
when they are in a semi-confluent state. In general, mammalian cells should be handled
gently. They should not be vortexed, vigorously pipetted or centrifuged at greater than
1500 g.

3. Cell feeding. Use prewarmed media and have cells out of the incubator for as little
time as possible. Use 10-15 ml forT-25's, 25-35 ml forT-75's and 50-60 ml forT-150's.
a. Suspension cultures. Feeding and subculturing suspension cultures are done
simultaneously. About every 2-3 days, dilute the cells into fresh media. The dilution you
use will depend on the density of the cells and how quickly they divide, which only you
can determine. Typically 1:4 to 1:20 dilutions are appropriate for most cell lines, b.
Adherent cells. About every 2-3 days, pour off old media from culture flasks and replace
with fresh media. Subculture cells as described below before confluency is reached.

4. Subculturing adherent cells. When adherent cells become semi-confluent, subculture
using 2 mM EDTA or trypsin/EDTA.

Trypsin-EDTA :

•a. Remove medium from culture dish and wash cells in a balanced salt solution

without Ca + + or Mg + + . Remove the wash solution,
•b. Add enough trypsin-EDTA solution to cover the bottom of the culture vessel and

then pour off the excess.

c. Place culture in the 37°C incubator for 2 minutes,
•d. Monitor cells under microscope. Cells are beginning to detach when they appear

rounded.
>*e. As soon as cells are in suspension, immediately add culture medium containing

serum. Wash cells once with serum containing medium and dilute as appropriate

(generally 4-20 fold).

EDTA alone:

•a. Prepare a 2 mM EDTA solution in a balanced salt solution (i.e., PBS without Ca ++ or
Mg ++ ).

b. Remove medium from culture vessel by aspiration and wash the monolayer to
remove all traces of serum. Remove salt solution by aspiration.

c. Dispense enough EDTA solution into culture vessels to completely cover the
monolayer of cells.

■d. The coated cells are allowed to incubate until cells detach from the surface.
Progress can be checked by examination with an inverted microscope. Cells can be

gently nudged by banging the side of the flask against the palm of the hand,
•e. Dilute cells with fresh medium and transfer to a sterile centrifuge tube,
•f. Spin cells down, remove supernatant, and resuspend in culture medium (or freezing

medium if cells are to be frozen). Dilute as appropriate into culture flasks.

5. Thawing frozen cells.

a. Remove cells from frozen storage and quickly thaw in a 37°C waterbath by gently
agitating vial.

b. As soon as the ice crystals melt, pipet gently into a culture flask containing
prewarmed growth medium.

•c. Log out cells in the "Liquid Nitrogen Freezer Log" Book.

6. Freezing cells.

•a. Harvest cells as usual and wash once with complete medium.

•b. Resuspend cells in complete medium and determine cell count/viability.

•c. Centrifuge and resuspend in ice-cold freezing medium: 90% calf serum/ 10% DMSO,

at 10 6 - 10 7 cells/ml. Keep cells on ice.
•d. Transfer 1 ml aliquots to freezer vials on ice.

■e. Place in a Mr. Frosty container that is at room temperature and that has sufficient

isopropanol.
•f. Place the Mr. Frosty in the -70°C freezer overnight. Note: Cells should be exposed

to freezing medium for as little time as possible prior to freezing
•g Next day, transfer to liquid nitrogen (DON'T FORGET) and log in the "Liquid Nitrogen

Freezer Log" Book.

7. Viable cell counts. USING A HEMOCYTOMETER TO DETERMINE TOTAL CELL

COUNTS AND VIABLE CELL NUMBERS (Reference: Sigma catalogue)Trypan blue is
one of several stains recommended for use in dye exclusion procedures for viable cell
counting. This method is based on the principle that live cells do not take up certain
dyes, whereas dead cells do.

1. Prepare a cell suspension, either directly from a cell culture or from a concentrated or
diluted suspension (depending on the cell density) and combine 20 pi of cells with 20 pi
of trypan blue suspension (0.4%). Mix thoroughly and allow to stand for 5-15 minutes.

2. With the cover slip in place, transfer a small amount of trypan blue-cell suspension to
both chambers of the hemocytometer by carefully touching the edge of the cover slip
with the pipette tip and allowing each chamber to fill by capillary action. Do not overfill
or underfill the chambers. 3. Starting with 1 chamber of the hemocytometer, count all
the cells in the 1 mm center square and four 1 mm corner square. Keep a separate
count of viable and non-viable cells. 4. If there are too many or too few cells to count,
repeat the procedure either concentrating or diluting the original suspension as
appropriate. 5. The circle indicates the approximate area covered at 100X microscope
magnification (10X ocular and 10X objective). Include cells on top and left touching
middle line. Do not count cells touching middle line at bottom and right. Count 4 corner
squares and middle square in both chambers and calculate the average. 6. Each large
square of the hemocytometer, with cover-slip in place, represents a total volume of 0.1
mm 3 or 10" 4 cm 3 . Since 1 cm 3 is equivalent to approximately 1 ml, the total number of
cells per ml will be determined using the following calculations:Cells/ml = average cell

count per square x dilution factor x 10 ;

Total cells = cells/ml x the original volume of fluid from which the cell sample was
removed; % Cell viability = total viable cells (unstained)/total cells x 100.

This Manual is still in the works so keep checking for updated versions.

Molecular Biology Laboratory Manual

Version 1,2

Denny R Randall
3/24/2009