buffer preparation

Lab 2 – Basic Techniques & ONs

Lab 2

Basic Techniques
Demonstration of sterile technique.
Lab 2A: Dilute 10X TE Buffer to Make 1X TE Buffer
Lab 2B: Determine the Concentration of an Unknown DNA Sample
Lab 2C: Streak out bacteria colonies

Buffers

Many of the experiments performed in molecular biology and biochemistry require
proteins to carry out a particular function, such as binding to DNA or cleaving a substrate. The
activity of these proteins is often very dependent on the pH, salt concentrations, and temperature
of the reaction mixture. In some cases a change of pH from 7.5 to 6.5 or a 10-degree change in
the temperature may cause greater than a 1000-fold reduction in the protein's activity. It is
therefore very important to understand the function of the proteins involved in each experiment
and know their optimal conditions for activity.

As little as 15 years ago, most enzymes used to manipulate DNA were difficult to obtain
because they were purified only by biochemists in research laboratories. In addition, there were
a very limited number of enzymes available. However, as more and more of the techniques in
molecular biology and biochemistry have become commonplace, companies have stepped in and
now provide these enzymes. Many of the techniques that we use in molecular biology and
biochemistry are now provided by these companies in the form of kits that include all enzymes,
reagents, buffers, protocols, and (frequently) controls for the experiment. These kits are often
very helpful, as well as convenient, for carrying out standard procedures. However, it is easy to
get very complacent about just following the instructions and not understanding what is actually
involved at each step of the protocol. You should understand enough about the procedure to
know the function of the kit's buffers, regardless of whether you have to personally make it. A
buffer is not a magic potion but a chemical solution containing a specific mixture of salts,
buffering agents, and sometimes reducing agents, detergents or cofactors, etc., in which each of
the components has a purpose and is included to optimize the reaction.

2-1


Solutions

Most of the solutions used in molecular biology and biochemistry are calculated on the
basis of the molarity of the solute. Sometimes a solution will be made on the basis of weight
percent, parts per million (ppm), or normality.

I. Molarity: A solution based on the number of moles of solute in a given volume of solution.

Molarity = moles of solute
liter of solution
Example 1: If 2.92 g of NaCl is dissolved in enough water to make 250 ml of solution,
what is molarity of the NaCl? (The molecular weight of NaCl is 58.5 g/mole.)

M = moles solute 2.92 g X 1 mole NaCl = 0.050 moles NaCl
liter of solution 58.5 g NaCl

= 0.050 moles
0.250 liters

= 0.2 M (or 200 mM)

Example 2: Calculate how much NaCl to use to make 300 ml of a 450 mM solution.
A 450 mM solution is 0.45 moles/ liter and 300 ml = 0.3 liter. (Remember to keep the
units constant in your calculations – forgetting to convert ml to liters or mg to g could
throw your calculations off 1000 fold).

0.45 moles X 0.3 liter X 58.5 g NaCl = 7.9 g NaCl
liter moles NaCl
Therefore, to make 300 ml of a 450 mM NaCl solution add 7.9 g NaCl to water. Once
the NaCl is dissolved, bring the final volume up to 300 ml by adding water. Why?

II. Percentage: A percentile of the total volume. There are two kinds of percentage solutions:
Percentage by weight (w/v), for dissolving solids, and percentage by volume (v/v), for diluting
concentrated % solutions.

% by Weight (w/v) = grams of solute
100 ml of solution

Example: How do you prepare a 50-ml solution of 20% (w/v) Sodium Dodecyl Sulfate
(SDS)?
20 g X 50 ml = 10 g SDS
100 ml

2-2


Therefore, to make 50 ml of a 20% (w/v) SDS solution, dissolve 10 g SDS in water and
bring the volume up to 50 ml with water.

% by Volume (v/v) = volume of concentrate X concentrate %
final volume of solution
Example: How do you prepare a 200 ml solution of 70% Ethanol from 100% Ethanol?

70% = ? X 100% 70% X 200 ml = ?
200 ml 100%

? = 140 ml of 100% Ethanol

Therefore, to prepare a 200 ml solution of 70% (v/v) Ethanol, start with 140 ml of 100%
Ethanol and add water (60 ml) to bring the final volume up to 200 ml.

The X Factor

In a molecular biology research lab, you will constantly need to make and use buffers. In
order to save time and space, molecular biologists often make concentrated stocks of solutions to
last over long periods of time. Such concentrated stocks take up less space. In addition, these
stocks are easily diluted for use when necessary. In this and other labs, you will often deal with
solutions that are labeled “10X,” “5X,” “100X,” etc. It is important to understand what this “X”
factor means.

The “X” factor simply indicates that the solution is in a concentrated form that must
usually be diluted to a “1X” concentration for use. For example, a 5X concentrated solution must
be diluted 5-fold, while a 100X concentrated solution must be diluted 100-fold. The dilutions
are usually done using water.

Dealing with the X factor eliminates the need to know the actual molar concentration of
the various components within the solution. You simply need to add water to make a 1X
solution. However, a good scientist should always understand the composition of his or her
reagents.

Example: Prepare 1 liter of 1X TBE buffer from a 10X TBE stock solution.

Below is a useful formula for doing dilution calculations:

V1C1 = V2C2

V1 = volume of stock buffer = ? V1C1 = V2C2
C1 = concentration of stock buffer = 10X ? (10X) = (1 liter) (1X)

2-3


V2 = volume of dilute buffer = 1 liter ? = (1 liter) (1X) / (10X)
C2 = concentration of dilute buffer = 1X ? = 0.1 liter

Therefore, to prepare 1 liter of 1X TBE from 10X TBE stock, you should add 100 ml of
10X TBE to 900 ml of water. (Note: Diluting a buffer or solution does not change its
gram or molar amount—only it’s concentration. The example above illustrates how a
small volume of a concentrated buffer (100 ml of 10X TBE) is equivalent to a large
volume of its diluted form (1 liter of 1X TBE).

Preparing Buffers

It is common in molecular biology to have to prepare complex buffers by diluting two or
more stock solutions. For example, if you want to prepare TE buffer (0.1 mM EDTA, 10 mM
Tris), it is best to simply dilute stocks of concentrated EDTA and Tris solutions.

Example: Prepare 100 ml of TE buffer from 0.5 M EDTA and 1 M Tris stocks.

Use the formula from above (V1C1 = V2C2) and solve for each of the two components
independently:

EDTA

V1 = volume of stock buffer = ?
C1 = concentration of stock buffer = 0.5 M (?) = (100 ml) (0.1 mM) / (500 mM)
V2 = volume of dilute buffer = 100 ml
C2 = concentration of dilute buffer = 0.1 mM (?) = 0.02 ml

Tris

V1 = volume of stock buffer = ?
C1 = concentration of stock buffer = 1 M (?) = (100 ml) (10 mM) / (1000 mM)
V2 = volume of dilute buffer = 100 ml
C2 = concentration of dilute buffer = 10 mM (?) = 1 ml

Therefore, to prepare 100 ml of TE Buffer from 0.5 M EDTA and 1 M Tris stocks, you
should add 0.2 ml of 0.5 M EDTA and 1 ml of 1 M Tris to 98.98 ml of water.

We strongly suggest that you do some of the buffer and dilution problems in the previous
quizzes. Similar problems will be on quizzes and exams in this class!

2-4


Lab 2A: Dilute 10X TE Buffer to Make 1X TE Buffer
(Each person in each group should make his/her own 1X TE)
1. Make up 25 ml of sterile 1X TE.
You will need to make 25 ml of 1X TE from a 10X TE stock and sterile water. Prepare this
in a sterile 50-ml tube. SAVE THIS SOLUTION. It will be used to dilute DNA in
experiment #3. To solve this dilution problem ask yourself “What volume of 10X TE stock
will I need in order to make 25 ml of 1X TE?” (See the example at the bottom of p. 2-3.)

V1 = ?
C1 = 10X TE
V2 = 25 ml
C2 = 1X TE
Answer the following questions. A course staff will check your answers.

1. How much 10X TE do you need to use? _____________

2. What pipetor will you need to dispense the liquid? __________________

3. How much sterile water do you need to use? _____________

4. What pipetor will you need to dispense the liquid? __________________

2-5


Lab 2B: Determine the Concentration of an Unknown DNA
Sample by comparison
Method A: Set up titration of DNA plus ethidium bromide and use the camera system to
estimate the concentration.

Many experiments in molecular biology involve manipulating small quantities of DNA to
digest, clone, or sequence. In working with small quantities of DNA it is often important to
know the concentration of the DNA samples in order to determine whether the experiment is
practical and feasible. Although in most cases the exact concentration is not critical, it is often
important to know the relative concentration of the DNA; i.e. whether you are working with 0.1,
1, 10 or even 100 micrograms (µg) of DNA. The following lab experiments are methods used to
determine the concentration of DNA in a sample. This exercise is not only useful for learning
how to measure the concentration of DNA, but it will also familiarize you with working with
pipetmen and the camera set-up, which will be used often in the course for a variety of
procedures.

Each person in each group will be given a different sample of plasmid DNA of unknown
concentration (Unknown DNA X or Unknown DNA Y) and a sample of plasmid DNA with a
known concentration (the DNA Standard). Each person in each group will determine the
concentration of their unknown sample by first serially diluting his/her DNA sample. Following
the addition of ethidium bromide, the DNA dilutions will be visualized using a UV light source.
The concentration of the Unknown DNA will also be measured by determining the amount of
UV light that it absorbs.

Ethidium bromide (EtBr) is a dye that intercalates between the DNA bases and emits
fluorescence. The amount of fluorescence emitted is directly proportional to the amount of DNA
in the sample (over a given range of DNA amount). The concentration of a sample of DNA can
therefore be determined by comparing the fluorescence of sample with that of a DNA sample of
known concentration. As little as 1-5 nanograms (ng) of DNA can be detected by this method.
The presence of any protein contaminants in a presumably pure DNA solution does not interfere
with the assay but there may be other contaminants in your DNA that may quench or possibly
contribute to the fluorescence.

Caution! Ethidium bromide, which is a potent mutagen, can intercalate into your DNA
as well as your sample DNA. Gloves should be worn when working with solutions of EtBr.
Prolonged exposure to UV light is also dangerous to your skin, and especially your eyes. To
minimize exposure make sure the UV light box is shielded and that you wear protective eye
goggles to efficiently block UV radiation.

2-6


Determine the concentration of DNA using ethidium bromide

Using ethidium bromide to measure the concentration of your DNA is a good method
when your DNA concentration is very low (<250 ng/µl) or if your sample is contaminated with
proteins or phenol.

In this experiment, each person will make a series of dilutions of a DNA sample of
known concentration (the DNA standard). Another series of dilutions will be made using
DNA samples of unknown concentration (one group partner gets Unknown DNA X and the
other partner gets Unknown DNA Y). Ethidium bromide will then be added to all DNA
dilutions, then the dilutions will be visualized under UV light. By comparing the fluorescence of
the unknown DNA samples with that of the known DNA samples (the standards), it should be
possible to estimate the concentration of DNA in the unknown sample.

1. Serially dilute your DNA sample (Unknown DNA X or Y) 5 times, by 5-fold dilutions.
a) Label 5 microcentrifuge tubes with “1X”, 2X, 3X, 4X, and 5X if you have the X sample or
1Y, 2Y, 3Y, 4Y and 5Y if you have the Y sample.
b) Using a P-200, aliquot 40 µl of TE into each of 5 labeled microfuge tubes.
Make sure that you have pipeted the correct volume into each tube by comparing it
with standard tube with the blue dye.
c) Use a P-20 to add 10 µl of the “Unknown DNA” (X or Y) to Tube #1 and vortex. (see the
figure on the next page). You have now diluted the DNA 5 –fold.
d) Add 10 µl of the 1X (or Y) to Tube #2 and vortex. You have now diluted the DNA 25 –
fold.
e) Add 10 µl of the 2X (or Y) to Tube #3 and vortex. You have now diluted the DNA 125 –
fold.
f) Add 10 µl of the 3X (or Y) to Tube #4 and vortex. You have now diluted the DNA 625 –
fold.
g) Add 10 µl of the 4X (or Y) to Tube #5 and vortex. You have now diluted the DNA 3125 –
fold.

2. Serially dilute your “DNA Standard” solution 5 times, by 5-fold dilutions.
a) Label 5 microcentrifuge tubes with “1S”, 2S, 3S, 4S, and 5S.
b) Using a P-200, aliquot 40 µl of TE into each of 5 labeled microfuge tubes.
c) Use a P-20 to add 10 µl of the ““DNA Standard”) to Tube #1S and vortex. (The
concentration of the DNA Standard is 1 µg/µl). (See the figure on the next page).
d) Add 10 µl of the 1S to Tube #2S and vortex.
e) Add 10 µl of the 2S to Tube #3S and vortex.
f) Add 10 µl of the 3S to Tube #4S and vortex.
g) Add 10 µl of the 4S to Tube #5S and vortex.

3. Pipet 40 µl of TE buffer into another tube to serve as a blank control.
You should now have 11 tubes in total — 5 dilutions the Unknown DNA, 5 dilutions of the
DNA Standard, and 1 with TE only.

2-7


Serial Dilutions for the EtBr Experiment

Label 5 tubes for diluting the unknown DNA sample (X or Y). Add 40 µl of 1X TE to each tube.

1 2 3 4 5

Add 10 µl of the unknown Transfer 10 µl of the sample in
DNA stock to the first tube. tube 1 to tube 2.

1

Unknown
DNA 1 2
Cap tube 2 and vortex to mix. (This DNA
Cap tube 1 and vortex to mix. (This DNA
has now been diluted 5-fold from the has now been diluted 25-fold from the
starting stock. )
starting stock. )

Transfer 10 µl of the sample in Transfer 10 µl of the sample in
tube 3 to tube 4. tube 2 to tube 3.

2
3

3
4 Cap tube 4 and vortex to mix. Cap tube 3 and vortex to mix. (This DNA
has now been diluted 125-fold from the
starting stock. )

Transfer 10 µl of the sample in
tube 4 to tube 5.

Repeat this process for the
known DNA standard.
4

5 Cap tube 5 and vortex to mix.

2-8


4. Prepare a 2 µg/ml solution of EtBr and spot 3 µl-drops of a diluted solution of EtBr
onto a petri dish as shown below.
a) Label two microfuge tubes “ET-200” and “ET-2”
b) Add 98 ul of TE to the “ET-200” tube and 198 µl TE to the “ET-2” tube.
c) Add 2 ul of 10 mg/ml EtBr stock to the ET-200 tube and vortex. This is a 1-50 dilution.
d) Add 2 ul of ET-200 EtBr stock to the ET-2 tube and vortex. This is a 1-100 dilution. (You
have now serially diluted your 10 mg/ml EtBr stock to a final concentration of 2 µg/ml.)
e) Place half of a clean 60-mm dish over the circular outline below. It will serve as a guide
on where to place your drops.
f) Add 3-µl drops of your 2 µg/ml EtBr solution such that there are two rows of 5 drops and
one drop off to the side. (See the figure below.)

Spots for
DNA standards
1 2 3 4 5 TE control

Spots for 1 2 3 4 5
unknown DNAs

5. To each of the above spots of 2 µg/ml EtBr place an equal volume (3 µl) of one DNA
sample dilution from the numbered tubes as shown and mix by pipeting up and down
several times. To the remaining spot add 3 µl of TE as a negative control.
This last spot is a zero DNA control and will indicate the background fluorescence of the
assay.

6. Being careful not to disturb the spots, take the plate to the dark room and photograph
the spots when exposed to UV light.
It is best to photograph your plate by placing it upside down on the UV light box. The
instructors or TAs will demonstrate how to use the camera set-up. You will use the camera
set-up throughout the course, so take notes on how to use it!!

7. Estimate the concentration of your unknown sample by comparing the intensity of the
spots with the spots of the DNA standards.
Later you will compare and comment on the calculated values of each method. Tape your
picture in your notebook neatly — use transparent tape. And label your picture with a pen or
thin marker.

2-9


Lab 2C: Streak out bacteria colonies

In last week’s lab you plated the bacteria library on selective media (LB-Kan) to select for cells
that were transformed with a plasmid from the library. We therefore need to perform a second
round of the screen to distinguish real mutants among the wild type cells. The best way to do
this is to streak out the colonies for single cells and examine the differences in growth of these
individual colonies.

A streak out is exactly like it sounds.

1. Take a sterile toothpick and gently touch one of
the colonies on the plate. A bacterial colony
typically has around 106 cells. Touching the
toothpick to the colony transfers a large number of Fig L2C.1 A streak out of the blue cell
the cells onto the toothpick. from the plate shows that there was a
mixture of cells on the toothpick. 1, 2,
2. Gently scrape the toothpick on a fresh plate. and 3 indicate the order of the streaks.
Normally this is just a short line (Streak 1 in Fig The streak out plate is shown after the
L2C-1). A large number of the cells will transfer cells have had a chance to grow ON.
from the toothpick to the plate. Often so many cells
will transfer that a patch of cells will appear after
the plate is incubated for a few days. This patch
may contain a number of cells from different
parents.

3. Take a new sterile toothpick and drag it in
perpendicular direction across the end of the first
streak and then extend it a short distance (Streak 2
in Fig L2B-1). The toothpick will pick up some Fig L2C.2 Examples of good and bad streak
cells from the first streak and then spread them on outs. The streak outs on the left and in the
the media as the toothpick is dragged across the middle are good because there are individual
plate. There may still be such a large number of colonies. The one on the right is bad because
cells that it forms a patch with a mixture of cells. it is a large patch with no individual colonies.
This patch may be a mixture of colonies.
4. Take a new sterile toothpick and drag it in Some of the colonies in the left and middle are
perpendicular direction across the end of second smeared because they were picked to make
streak and then extend the streak in a zig-zag overnight cultures.
fashion across a section of the plate (Streak 3 in Fig
L2B-1). The toothpick will pick up a few cells from the second streak and then spread them on
the media as the toothpick is dragged down the plate. There will now be so few cells on the
toothpick that the individual cells will deposited on the plate a disperse manner. When these
cells grow they will form individual colonies instead of a patch of cells that came from different
parents. All the cells in a colony will be generated from one cell and so they will be clonal. The
example shows that the streak out contained a mixture of blue and white cells. However,

2-10


individual colonies at the end of the streak out are clonally pure. Examples of good and bad
streak outs are shown in Fig L2C.2.

We are going to use the streak outs not only to make sure that the cells are clonally pure but to
also rescreen the growth on Kan.

Protocol for streak outs of colonies from the library plating.
1. Each student will be given 1 LB-Kan (two black lines on side) plate.
a) Using a ruler and a fine point Sharpie draw two perpendicular lines on the back of
each plate to divide it up into quarters (see Fig L2C.1).
b) Label each quadrant with
(a) Your Class day (e.g. T- Tuesday lab, W- Wednesday lab, H- Thursday lab)
(b) Your Group Number (e.g. 1, 2, 3 etc)
(c) Your initials (e.g. AV, JM, etc)
(d) The numbers of the clone (e.g. 1, 2, 3 or 4)
(e) The year (e.g. .10)
For example if you are a member of group 2 in the Thursday and your initials are AB, then
you should label the four quadrants: H2AB1.10, H2AB2.10, H2AB3.10 and H2AB4.10.

Do not write on the lids! Lids can be switched, causing strains to be mixed up.

2. Select a colony from the LB-Kan library transformation plate and streak it out onto the
first quadrant on the plate. Remember that the plate is flipped so the quadrant will be the
opposite side!!!
a) As described above, use one toothpick to pick the colony from the transformation
plate and make a short streak on one quadrant on the plate.
c) Use a fresh toothpick to make a short diagonal streak across the previous streak on
the plate.
d) Use another fresh toothpick to make one streak across the second streak and then
zig-zag the streak several times.

3. Repeat the streak out process picking 3 different colonies from the library plate that you
made last week. You should mark the back of the library plate which colonies that you picked.

4. These plates will be incubated at 37°C for several days until the streak outs are well
developed and single colonies are visible.

2-11

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