Download Bruce Wallace Biotechnology Lab Program Student Guide 5th

Bruce Wallace
Biotechnology Lab Program
Student Guide
5th Edition
PREFACE:
The significance of the science you are about to do is sometimes taken for granted as the protocols have been
worked and reworked so that there is a high probability that you will succeed at producing the desired molecular
product. The work that you are about to do is based on Nobel Prize–winning science. Werner Arbor, Daniel
Nathans and Hamilton Smith received the Nobel Prize for their work with restriction enzymes. Stanley Cohen,
Paul Berg and Herb Boyer received the Prize for making the first recombinant DNA molecule. The recombinant
DNA molecule that you are about to use extends beyond their work as it uses a gene from a eukaryote rather
than prokaryotic organism. As recently as 1993, Kary Mullis received the Nobel Prize for his discovery of the
Polymerase Chain Reaction, some elegant chemistry that you will be using in Laboratory 8. So the science you
will be covering over the next few weeks is significant and will continue to play an important role in the
development of biotechnology and medicine.
Your teacher deserves a great deal of credit for making this laboratory experience possible. Although Amgen
provides the equipment and the supplies needed to implement the labs, your teacher has provided many hours of
preparation time, often involving weekends and evenings, to make them happen. If you’ve enjoyed this
laboratory experience, please remember to thank your teacher for making it a reality.
This educational outreach program is largely the result of the efforts of Dr. Bruce Wallace, an Amgen scientist
who strongly believed that the biotechnology industry had a responsibility to contribute to the science education
of our society. Before Dr. Wallace’s untimely passing, he was able to see his educational outreach program
grow and evolve into the adventure of discovery upon which you are about to embark.
We are able to bring this program to your school because of several key partnerships: Amgen Foundation,
Foundation for Pierce College, Los Angeles/Orange County Biotechnology Center, The Bio-Bridge Institute
(biobridge.ucsd.edu/a1), New England Biolabs, Fotodyne, Invitrogen, Rainin Pipettes, VWR and Bio-Rad.
Should you have any questions about these laboratories, please feel to e-mail me at the address below.
Martin Ikkanda
Professor of Biology
Los Angeles Pierce College
[email protected]
TABLE OF CONTENTS
Lab number
Pages
Lab title
An Introduction to Microvolumetrics
and Pipetting
Restriction Analysis of
pARA and pKAN-R
pARA-R Restriction Digest:
An Introduction to Plasmids and Restriction Enzymes
Ligation of pARA/pKAN-R Restriction Fragments
Producing a Recombinant Plasmid, pARA-R
Confirmation of Restriction and Ligation Using
Agarose-Gel Electrophoresis
Confirmation of pARA-R Restiction Digest
Transforming Escherichia coli with a Recombinant Plasmid
Transformation of Escherichia coli
with pARA-R
Preparing an Overnight Culture of Escherichia coli
Purification of mFP from
an Overnight Culture
Genomic DNA Extraction From Buccal Epithelial Cells
Lab manual design by Lucy Reading
Cover illustration courtesy of Ken Eward
Lab1:
An Introduction to Microvolumes and Pipetting
The purpose of this laboratory is to provide you with a hands-on experience using some of the important tools
and techniques commonly used in molecular biology and introduce you to some of the volumetric
measurements that are most often used in this field of science. The laboratory will provide you with an
opportunity to practice some of the skills you will need to build a recombinant DNA molecule. The instruments
and supplies that you will be using over the next few weeks are identical to the ones that are used in research
laboratories.
While the theoretical foundations upon which biotechnology and DNA sciences have been built extend back to
the early 1900s, most of the laboratory techniques utilized are relatively recent. And though the techniques you
will be learning over the next few weeks have become routine in modern research laboratories, few high school
and college students have an opportunity to do such sophisticated molecular biology.
If and when you take a chemistry class, one of the things you will quickly notice is the differences in the
quantities of reagents and chemicals that you use. In a typical chemistry lab, volumes are measured in large
graduated cylinders. Solutions are often measured in 50, 100 or 200 milliliters (mL) volumes. Weights of solids
are generally expressed in grams (g). In the molecular biology lab, volumes are frequently measured in
microliters (µL); 1 µL is equal to 0.001 mL. Weights are often expressed in terms of micrograms (µg) or
nanograms (ng); 1 µg is equal to 0.000001 gram and 1 ng is equal to 0.000000001 gram.
You might be wondering why molecular biologists use such small volumes and amounts of materials. The
reason is related to the cost of these materials and the difficulty involved with obtaining them. For example, you
will be given some specially engineered plasmids (DNA) in the next laboratory. If this DNA were sold “by the
pound,” it would cost around $360,000,000 per pound. So don’t be surprised if we only give you a tiny amount
of these DNA molecules. The reason why these chemicals are so expensive is related to the difficulty in
preparing them in pure form. Many of these chemicals are produced within living organisms, such as bacteria,
and have to be purified and separated from all of the other thousands of substances in the cell. Molecular
biology, however, really requires this level of purity and precision. As you do this lab work, keep in mind that
you are doing real-world molecular biology.
Materials
Reagent
Solution 1
Solution 2
Solution 3
Distilled H2O (dH2O)
0.8%
Agarose gel (pre-made)
1 x SB (or 0.5x TBE)
Equipment & supplies
1.5 mL microfuge tubes
P-20 micropipettor (2-20 µL)
Disposable pipette tips
Permanent marker
Electrophoresis equipment
Power supply
Plastic microfuge tube rack
Methods
The Digital Micropipette
Molecular biology protocols require the use of adjustable micropipettes. Micropipettes are used to dispense
different volumes of liquids. While researchers will have several kinds of micropipettes at their lab bench, these
laboratories have been designed to utilize a P-20. The P-20 is engineered to dispense liquid volumes between 2
and 20 µL. This is a high-quality, precision instrument, and it is essential that you learn to use it properly.
Please read and follow these precautions:
- Do not set the adjustment below 2 µL or above 20 µL unless instructed to do so by your teacher.
- Do not use the micropipette without the proper disposable tip firmly attached to the barrel. Failure to use a
pipette tip will contaminate the pipette barrel.
- Do not lay down a micropipette with fluid in the tip or hold it with the tip pointed upward. If the disposable
tip is not firmly seated onto the barrel, fluid could leak back into the pipette.
- Avoid letting the plunger “snap” back when withdrawing or ejecting fluid; it will eventually destroy the
piston.
--------------------------------------------------------------------------------------------------------------------------------------- When aspirating (drawing up) a solution, push the plunger to the first stop and lower the pipette tip below the
level of the solution that you are sampling. You should be holding the tube containing the solution in your hand
about eye level. It’s important to actually see the solution enter the pipette tip.
- Slowly release the plunger and allow the liquid to move into the pipette tip. Be certain that you’re not
aspirating air into the tip.
- When dispensing (pushing out) the liquid, place the pipette tip into the tube that will receive the solution.
Position the tip so that it touches the side and near the bottom of the tube. Slowly push down on the plunger to
the first stop and then to the second stop. Keep your thumb on the plunger and remove the tip from the tube into
which you’re dispensing the liquid. This will avoid reaspirating the liquid into the pipette tip. Be certain that
you see the solution leaving the tip.
- Remove the tip by ejecting it into a waste container; there is an eject button on the pipette. If you’re
dispensing the same reagent into separate tubes and there is no danger of cross contamination, you can use the
same tip several times. To avoid contamination, it is good practice to deposit each reagent on to the sidewall
near the bottom of the microfuge tube without touching any of the other reagents. This technique allows you to
use the same tip to dispense a reagent into several tubes that contain a different reagent.
- When dispensing a new reagent, always use a fresh tip to avoid contamination.
Pipetting Exercise 1
1. Find the display window on the handle of the micropipette and note its setting. Turn the knurled knob in the
handle clockwise to decrease the volume or counterclockwise to increase the volume. Turning this knob
changes the distance the plunger will travel. The figures below represent some pipette settings and the volumes
of liquid dispensed.
2. Place a disposable tip onto the end of the pipette barrel. Using your thumb and index finger in a twisting
motion, check to see that the tip is firmly seated onto the barrel. Avoid touching the pointed end, as this may
contaminate the tip.
Remember that you must have a tip in place when using the pipette.
3. Place your thumb on the button that activates the plunger. Push down on this button with your thumb and
notice that it has a “stop” position. If you exert a little more pressure with your thumb, you can push the button
of the plunger to a second stop. The second stop pushes a small volume of air into the tip to eject the solution.
Pipetting Exercise 2
1. Use a permanent marker to label three reaction tubes A, B and C.
2. The table on page 1.4 summarizes the contents of each tube, but follow the directions that begin with step 3
to set up the samples.
3. Set the P-20 micropipette to 2 µL and dispense dH2O into tubes A, B and C.
4. Eject the tip into the plastic waste container and replace with a fresh tip.
5. Place 4 µL of solution 1 into tube A.
6. Eject the tip into the plastic waste container and replace with a fresh tip.
7. Use a fresh tip and dispense 4 µL of solution 3 into tube A.
8. Use a fresh tip and dispense 8 µL of solution 2 into tube B.
9. Use a fresh tip and dispense 8 µL of solution 3 into tube C.
10. Save all three tubes for the next part of the lab.
Checking the Accuracy and Consistency of Pipetting
1. Tubes A, B and C should each contain 10 µL of solution.
2. Set your P-20 micropipette to 10 µL and place a fresh tip onto the barrel.
3. Carefully check the volume of each of microfuge tube. There should be 10 µL in each of these tubes.
4. Save tubes A, B and C for the next part of the lab.
Using Gel Electrophoresis to Separate Molecules
Gel electrophoresis is a method that uses an electrical current and a gel matrix (meshwork) to separate
molecules such as DNA and proteins. The molecules that are being separated are either negatively charged or
are made to be negatively charged. Using an electrical current, the charged molecules are then forced through a
meshwork of material that will sort out the molecules according to their sizes, although molecular shape and
degree of electro-negativity will influence movement through the gel. Because the molecules are negatively
charged, they will migrate through the gel toward the positive (red) electrode. The more negatively charged, the
faster the molecule will migrate.
In this laboratory, your teacher has made a gel composed of agarose, a polysaccharide (complex sugar). The
agarose is mixed with an electrolytic solution called Sodium Borate (SB). This solution contains ions, which are
electrically charged atoms. These ions help conduct the electrical current through the gel. As the molecules are
drawn toward the positive electrode, the smaller molecules are able to move in and around this agarose network
much more quickly than the larger molecules. Thus, over the length of the gel, the molecules become separated
by size.
1. Your teacher has already prepared an agarose gel for you, but you will need to cover the agarose gel with
the appropriate amount of SB buffer to run the gel properly. Two groups will share each gel. Take the box to
the power supply you will use to run the gel.
2. Check to make certain that the gel is positioned in the gel box so that the “wells” of the gel are located
toward the negative (black) electrode. The dyes are negatively charged and they will move toward the
positive (red) electrode.
3. Fill the box with 1x SB buffer (there are several plastic containers containing this buffer in the lab) to a
level that just covers the entire surface of the gel to a depth of 1–2 mm. Check to see that the gel is covered
with buffer and that no “dimples” appear over the wells; add more buffer if needed.
4. Set the micropipette to 10 µL and load each sample into a separate well as indicated by your teacher. Use a
fresh tip for each sample. Remember that your group will be sharing this gel. One group will load their
samples in three wells on the left while the other group will use the three wells on the right. You may wish to
record which solution you place in each well.
5. When loading each sample, center the pipette tip over the
well and gently depress the pipette plunger to slowly expel
the sample. Use your other hand to support your pipette hand
to avoid shaking. Because their densities are greater than the
SB buffer, the dyes will sink into the wells.
6. Close the cover tightly over the electrophoresis chamber. Connect the electrical leads to the power supply.
Be certain that both leads are connected to the same channel with the cathode (–) to cathode (black to black)
and anode (+) to anode (red to red).
7. Turn on the power supply and set the voltage to 130–135 v.
8. After two or three minutes, look at the dyes to make certain they are moving toward the positive (red)
electrode. You should begin to see the purple dye (called Bromophenol blue) beginning to separate from the
blue dye (Xylene cyanole).
9. In approximately 10 minutes, or when you can distinguish all three dyes, turn off the power switch and
unplug the electrodes from the power supply. Do this by grasping the plug at the power supply—not by
yanking on the cord. Carefully remove the cover from the gel box so that you can better see the dyes in the
gel.
10. On a piece of notebook paper, record the banding or color pattern in each of the lanes containing your
samples. Use this information to answer the questions in the “Conclusions.”
11. Leave the gels in the gel box.
Conclusions
1a. The dyes that you separated using gel electrophoresis were:
Orange G (yellow), Bromophenol blue (purple) and Xylene cyanole (blue).
What electrical charge did these dyes carry?
1b. What evidence allowed you to arrive at this conclusion?
2a. Molecular size can play a role in separation with small molecules moving through the gel matrix more
rapidly than larger molecules. The formula (or molecular) weights for these dyes are Orange G (452.38),
Bromophenol blue (669.98) and Xylene cyanole (538.62). From your results, did it appear that these molecules
were separated clearly on the basis of size?
2b. What other factors may have played a role in the separation of these dyes?
2c. Which tube contained a single dye? A, B or C?
2d. Name this dye.
3. When aspirating a solution, why is it important to actually see the solution enter the pipette tip?
4a. After loading your gel, did any solution remain in tubes A, B or C?
4b. What could account for solution remaining in these tubes?
Lab2:
Restriction Analysis of pARA and pKAN-R
Plasmids are circular pieces of DNA that are found naturally in bacterial cells. The plasmids used in
molecular biology have been modified through genetic engineering to facilitate gene cloning and protein
production (gene expression) in bacteria. Antibiotic resistant genes have been engineered into these plasmids
and function as selectable markers—that is to say, these genes allow us to select between bacteria that harbor
the plasmids from those that do not. If a bacterium carries a plasmid with an antibiotic resistant gene, the
bacterium will be able to grow and reproduce in the presence of that antibiotic; those bacteria without the
plasmid will not be able to grow. Thus, antibiotics can be used to select bacteria that are resistant and
presumably carry a plasmid with the resistant gene from those bacteria that do not carry the plasmid. Two
plasmids will be used in this laboratory: pARA contains a gene for ampicillin resistance, ampr, and pKAN-R
contains a gene for kanamycin resistance, kanr.
The purpose of this laboratory is threefold: 1) to introduce a method commonly used to analyze the genetic
elements of plasmid DNA; 2) to examine the role and nature of restriction enzymes; and 3) to take the first steps
in producing a recombinant DNA molecule.
The plasmid pARA is 4058 base pairs (bp) in size. A “base pair” would be adenine:thymine or
guanine:cytosine and is the common method used to express the size of DNA molecules. The plasmid carries
the ampr gene, which encodes the protein beta lactamase, an enzyme that destroys the antibiotic ampicillin.
Beta lactamase, then, enables bacteria to reproduce in the presence of ampicillin. In addition, pARA carries a
gene for the AraC protein, a protein that helps the bacterium make proteins encoded by genes inserted into this
plasmid. A gene, even a foreign one, can be expressed (produced) if it is inserted into a specific location in this
plasmid. The region of pARA labeled pBAD, in the plasmid map, indicates the site where RNA polymerase
needs to bind to initiate transcription. The sites labeled “BamH I” and “Hind III” represents restriction sites for
these two restriction enzymes. Study the plasmid map below and locate these plasmid components.
The plasmid pKAN-R carries the kanamycin resistant gene, kanr, which encodes a phosphotransferase, an
enzyme that transfers a phosphate group to the kanamycin molecule destroying its antibiotic effects. Kanamycin
is an antibiotic that kills bacteria by preventing them from making proteins. If a cell cannot synthesize proteins,
it will die. The kanr gene confers resistance to kanamycin for bacteria that have taken up this gene. In addition
to kanr, the plasmid carries the gene for mutated Fluorescent Protein, mFP, called red fluroescent protein or
“rfp.” The pKAN-R plasmid is approximately 5,408 bp in size.
The fluorescent protein gene was originally isolated from Discosoma sp, a sea anemone found in the IndoPacific ocean. The wild-type gene has been mutated, through a process called directed evolution, to produce
colors that are several times brighter than the wild-type protein. The term “wild type” refers to the original gene,
the one that you would find in nature. The rfp gene has been engineered into the plasmid pKAN-R. Note that
the rfp gene for mFP has both BamH I and Hind III restriction sites on either side. A “restriction site” marks the
specific location where an enzyme will cut the DNA plasmid. If pKAN-R is digested with BamH I and Hind III,
the rfp gene will be physically cut from the plasmid. During this laboratory, then, you will remove the rfp gene
from pKAN-R and remove the small 40bp fragment from the pARA plasmid using the same enzymes. During
the next laboratory, you will insert the rfp gene into pARA producing a recombinant DNA molecule.
Materials
Reagent
pARA (80 ng/ µL)
pKAN-R (80 ng/ µL)
Restriction enzymes
(BamH I + Hind III)
2.5x restriction buffer
Distilled water, dH2O
Equipment & supplies
P-20 Micropipette And Tips
1.5 Ml Microfuge Tubes
Minicentrifuge
37°C Water Bath
Permanent Marker
Methods
Preparing the pARA-R restriction digest
This laboratory protocol uses the restriction enzymes BamH I and Hind III to digest the plasmids pARA
and pKAN-R. This is the first step in making a recombinant DNA molecule.
1. Obtain the following four microfuge tubes: pARA, pKAN-R,
BamH I and Hind III (enzyme mix) and 2.5x buffer.
2. Obtain four clean 1.5 mL microfuge tubes and use a
marker to label a set of four 1.5 tubes as follows:
A + = pARA + BamH I and Hind III
A - = undigested pARA (pARA without enzyme)
K+ = pKAN-R+ BamH I and Hind III
K- = undigested pKAN-R(pKAN-R without enzyme)
3. The reaction matrix summarizes the reagents used in the restriction digest.
To set up the digest, follow the specific directions beginning at step 4.
4. Use a fresh tip and add 4µL of 2.5x restriction buffer to all four tubes.
5. Add 2µL of dH2O to tubes labeled A- and K -.
What is the purpose of this step?
6. Use a fresh tip and add 4µL of pARA to tubes labeled A+ and A-.
7. Use a fresh tip and add 4µL of pKAN-R to tubes labeled K + and K -.
8. Add 2µL of enzyme mix, containing BamH I and Hind III, to the A+ and K + tubes.
Add the enzymes directly into the solution at the bottom of the microfuge tube. Be certain to use a new tip for
each tube to avoid contamination. After the addition of the enzymes, gently pump the solution in and out with the
pipette to mix the reagents and cap the tubes .
9. If there is a minicentrifuge available, set the tubes into the rotor, being certain the tubes are in a balanced
configuration, and spin the tubes for four seconds. This brief spin will pool all of the reagents at the bottom of
each tube.
10. Place all four tubes into the 37°C water bath, and incubate for at least 60 minutes.
Following the 60 minute incubation, the digest can be kept frozen, at
–20°C, until time is available for electrophoresis.
Conclusions
Review the restriction maps of plasmids, pARA and pKAN-R. Because BamH I and Hind III are specific
restriction endonucleases, they will consistently cut DNA wherever it encounters the six-base recognition
sequences indicated below. The precise location that is cut is called a restriction site. The DNA molecule
consists of two strands of nucleotide building blocks. These building blocks are oriented in the opposite
direction on each strand. Thus, the two stands that makeup a DNA molecule are said to be “anti-parallel.” For
convenience, we can say that one strand is oriented in a 5’ (“five prime”) to 3’ (“three prime”) direction while
the other strand is oriented 3’ to 5’. Careful examination of the restriction sequences will reveal that the
sequence of nucleotides is a palindrome; that is to say, it reads the same on both strands when read in a 5’ 3’
direction.
Therefore, whenever Hind III encounters this six-base sequence, it will cut the DNA helix between the adjacent
adenine bases. This leaves four unpaired bases forming a “sticky end.”
1a. What is the recognition sequence for BamH I?
1b. In a 5’ 3’ direction, what sequence of bases represents the “sticky-ends for each?”
2a. Examine the pARA and pKAN-R plasmid maps and fill-in the following:
pARA digestion will yield
fragments and will be
base pairs in length.
pKAN-R digestion will yield
fragments and will be
bp and
bp in length.
2b. Assume you were given a culture of bacteria carrying one or both of these plasmids.
3. Design a simple experiment that you could use to determine which of these plasmids,
pARA or pKAN-R, the bacteria in the culture were carrying.
Lab2a:
pARA-R Restriction Digest:
An Introduction to Plasmids and Restriction Enzymes
Two powerful but fundamental tools used in biotechnology are restriction enzymes and bacterial
plasmids. Restriction enzymes allow molecular biologists to cut DNA molecules from different organisms and
recombine the molecular pieces to produce recombinant DNA molecules. Plasmids are circular pieces of DNA
that are naturally found in bacteria. Through recombinant DNA technology and restriction enzymes,
recombinant DNA plasmids can be engineered to clone genes or to express proteins encoded by genes.
Restriction enzymes were first observed by Werner Arbor in 1962. Arbor discovered that some bacteria
appeared to use a primitive immune system that prevented viral DNA from replicating within the infected host
cell. Some years later, it was revealed that this immune mechanism involved a class of proteins now known as
restriction enzymes. The name is derived from the enzyme’s ability to restrict the growth of viruses in the
bacterial cells. Restriction enzymes accomplish this by breaking a bond in the sugar-phosphate backbone of the
viral DNA- the enzymes cut the viral DNA into small fragments.
The restriction enzymes that were first identified appeared to digest the DNA molecule randomly. Later,
restriction enzymes were found and purified that would cut the sugar-phosphate backbone at a specific location
or within a specific nucleotide sequence, commonly four to six nucleotides in length. Table 1 identifies some of
these specific restriction enzymes, their source and the nucleotide sequences each recognizes. In 1978, Daniel
Nathans (Johns Hopkins University), Hamilton Smith (Johns Hopkins University) and Werner Arbor received
the Nobel Prize for Medicine for their work with restriction enzymes.
Table 1.
Restriction enzymes used in this laboratory.
-- indicate sites where the sugar-phosphate backbone is cut or cleaved.
When restriction enzymes cut or digest DNA, the fragments that result—called restriction fragments—have
several unpaired bases extending from their cut ends. These are called “sticky ends.” If DNA molecules from
different sources are digested using the same restriction enzyme, the unpaired bases from each piece should be
able to join (or anneal) together as the unpaired bases at the sticky ends will be complementary—A:T and G:C.
It is this unique attribute of restriction enzymes that enable genetic engineers to combine DNA fragments from
different organisms to produce recombinant DNA molecules.
Figure 1.
(a) DNA molecule with BamH I and Hind III restriction sites (bold). The arrows indicate sites where enzymes
will cut the sugar-phosphate backbone of the DNA molecule. (b) The lower DNA molecule indicates the
location of the “sticky ends” (bold).
Bacterial plasmids are relatively small, circular pieces of DNA that bacteria can carry in addition to their
genomic DNA (single chromosome). In nature, the plasmid DNA frequently carries one to several genes that
help the bacterium survive—perhaps by providing resistance to an antibiotic. Bacteria can pass along plasmids
during conjugation (mating). The bacteria we use in the laboratory have been mutated, so they cannot exchange
plasmids during sexual reproduction.
Naturally occurring plasmids have been engineered to perform specific functions: typically, gene cloning and
gene expression. This laboratory examines pARA-R, a recombinant DNA plasmid that has been engineered to
express the rfp gene to produce a mutant Red Fluorescent Protein (mFP). The plasmid contains various control
elements that allow a bacterium carrying this plasmid to express this foreign gene. The gene was originally
obtained from the genome of Discosoma sp., a sea anemone from the Indo-Pacific Ocean. The plasmid map
below indicates some of the important control regions, araC and PBAD, and the location of the rfp gene. In
addition, the map indicates the location of two restriction sites: one for Hind III and one for BamH I. How might
you go about cutting out the rfp gene? Also note, the plasmid carries an antibiotic resistance gene, ampr. This
gene will enable a bacterium carrying this plasmid to live in an environment containing the antibiotic
ampicillin.
Materials
Reagent
Para (70 Ng/ µL)
Restriction Enzymes (Bamh I + Hind Iii)
2.5x Restriction Buffer
Distilled Water (dH2O)
Equipment & supplies
P-20 Micropipette and Tips
1.5 mL Microfuge Tubes
Minicentrifuge
37°C Water Bath
Permanent Marker
Methods
Restriction Digest of pARA-R
The purpose of this laboratory is twofold: 1) to examine the role of restriction enzymes and their importance in
genetic engineering; 2) to examine a bacterial plasmid and how it is used in biotechnology.
This laboratory protocol uses the restriction enzymes BamHI and Hind III to digest the recombinant plasmid,
pARA-R. The restriction digest will isolate from pARA the rfp gene from the larger fragment of the plasmid
that containing ampr, araC and PBAD. The protocol uses a control, undigested pARA-R, along with a DNA
size marker or ladder that will help you identify and confirm the sizes of the restriction fragments.
Preparing the pARA-R Restriction Digest
1. Obtain the following three 1.5 mL microfuge tubes from your teacher:
pARA-R, enzyme mix and 2.5x restriction buffer.
2. Obtain two clean 1.5 mL microfuge tubes and use a marker to label the tubes as follows: “A+” and “A-.”
Include your group number and class period on each tube, so that you can locate them for the next lab
period.
3. The reaction matrix summarizes the reagents used in the restriction digest. To set up the digest, follow the
specific directions beginning at step 4.
4. Use a fresh tip and add 4µL of 2.5x restriction buffer to both tubes.
5. Add 2µL of dH2O to tube labeled A-.
What is the purpose of this step?
6. Use a fresh tip and add 4µL of pARA-R to tubes labeled A+ and A-.
7. Bring the A+ tube to your teacher, who will dispense the enzyme mix into the tube, or if you were given
this enzyme mix, carefully add 2µL of the enzyme mix directly into the solution in tube A+ containing
plasmid and buffer. After the addition of the enzymes, cap the tube and gently flick the lower portion of each
tube to mix the contents.
8. If there is a minifuge available, set the tubes into the rotor, being certain the tubes are in a balanced
configuration, and spin the tubes for four seconds. This brief spin will pool all of the reagents at the bottom of
each tube.
9. Place both tubes into the 37°C water bath, and incubate for at least 60 minutes.
10. Following the 60-minute incubation, your teacher may place the tubes into the freezer until you are ready for
electrophoresis (Lab 4a).
Digested plasmids can be kept at -20°C indefinitely.
Conclusions
Review the restriction map of the pARA-R plasmid. BamH I and Hind III are specific restriction enzymes and
will consistently cut the double-stranded DNA wherever they encounter their respective six-base recognition
sequence given in the table on page 2a.1. These locations cut are called restriction sites. The DNA molecule
consists of two strands of nucleotide building blocks. These building blocks are oriented in the opposite
direction on each strand. Thus, the two stands that makeup a DNA molecule are said to be “anti-parallel.” For
convenience, we can say that one strand in oriented in a 5’ (“five prime”) to 3’ (“three prime”) direction while
the other strand is oriented 3’ to 5’. Careful examination of the BamH I and Hind III restriction sequences will
reveal that the nucleotide sequences are palindromes; that is to say, they read the same on both strands when
read in a 5’ • 3’ direction.
Therefore, whenever Hind III encounters this six-base sequence, it will cut the DNA helix between the adjacent
adenine bases. This leaves four unpaired bases forming a “sticky end.”
1a. What are the recognition sequences for Hind III and for BamH I?
1b. In a 5’ 3’ direction, what sequence of bases represents the “sticky-ends?”
2a. Examine the pARA-R plasmid map and fill in the following:
How many restriction fragments will result from the digestion of pARA
with BamH I and Hind III?
2b. What will be the approximate lengths, in base pairs, of these restriction fragments?
2c. Which restriction fragment will carry the ampr gene?
2d. Which restriction fragment will carry the rfp gene?
3. Assume your teacher gave you a culture of bacteria. The culture could be one containing bacteria carrying the
plasmid pARA-R or a culture containing bacteria without the plasmid. Design a simple experiment that you
could use to determine which of these cultures you were given.
Lab3:
Ligation of pARA/pKAN-R Restriction Fragments Producing A Recombinant Plasmid, pARA-R
In this laboratory the restriction fragments produced during Lab 2 will be ligated, or bonded together, using
DNA ligase, making new recombinant plasmids. These newly formed plasmids will represent recombinant DNA
molecules because the four restriction fragments have been recombined in different ways to produce new
constructs. For example, assume that the four plasmid fragments were represented by the letter A, A’, K and R,
where A and A’ represent the pARA fragments and K and R represent the two fragments resulting from the pKANR digest. Plasmids could be represented by any combination of two letters, such as AK or A’R, and any
combination of even numbered fragments, such as AKA’R or ARAAKK and so forth. As you can see, there are
many kinds of recombinant molecules that could result from mixing together these restriction fragments.
As you will remember, the restriction enzymes we are using are BamH I and Hind III. Cutting the plasmids at the
BamH I and Hind III restriction sites leave “sticky ends.” The sticky ends on the cut DNA can be ligated to any
other fragment of DNA with a complementary sticky end. Examine the pARA plasmid map, below, to see the
locations of the BamH I and Hind III restriction sites and the sticky ends that form on the 5’-ends of its restriction
fragment.
Because pARA has one BamH I and one Hind III restriction site, the digest will leave two fragments. The
restriction fragments are depicted below. It is important to remember that the large restriction fragment carries the
ampr gene, the gene that provides resistance to ampicillin. The smaller fragment does not carry any genes.
The plasmid pKAN-R has one BamH I and one Hind III restriction site that flank the rfp gene. The digestion
of pKAN-R will leave two fragments, one will be 4706 bp and the other will be 702 bp.
Ligation will bond any two BamH I sticky ends together and any two Hind III sticky ends together. You
should be able to see that many different combinations of fragments are possible. The combination of interest to
us is the 4018bp pARA fragment recombined (containing the ampr gene) with the 702bp pKAN-R fragment
(rfp gene). The combination of these two fragments will yield a recombinant plasmid we will call pARA-R.
The ligation of the 702bp pKAN-R fragment will place the rfp gene into the plasmid at a location that will
allow a bacterium to synthesize (express) the mutant Fluorescent Protein, mFP.
The restriction fragments are initially held together by the hydrogen bonding between the nucleotide
bases that makeup the sticky ends. You may recall that adenine and thymine share two hydrogen bonds
while cytosine and guanine share three. This helps to ensure that only complementary sticky ends will
match up.
Hydrogen bonds are weak chemical bonds, and they are inadequate to hold the sticky ends together
permanently. The enzyme DNA ligase, with energy supplied by ATP, will form covalent bonds between the
sugar and phosphate groups of the DNA backbone. In the diagram below, you can see the positions of these
bonds on each side of the DNA molecule. When the covalent bonds are formed, the bonds complete the
phosphodiester linkage between the two sugars and the phosphate group on each strand. The resulting
chemical bonds are a relatively strong bond.
Materials
Reagent
Digested pARA (A+ from lab 2)
Digested pKAN-R (K+ from lab 2)
5x Ligation buffer with ATP
T4 DNA ligase in “lig” tube
Distilled water
Equipment & supplies
P-20 Micropipette and Tips
70°C Water Bath
Plastic Microfuge Tube Rack
Permanent Marker
Methods
1. Obtain your A+ and K+ tubes from the rack at the front of the class. Place the two tubes in the 70°C
water bath for 30 minutes. This heat exposure will denature (inactivate) any BamH I and Hind III that might
be active.
Why is this important?
2. While your tubes are in the water bath, obtain the 5x buffer and a Ligase tube from the instructor. The
ligase tube contains 2µL of DNA ligase. Label this tube with your initials.
3. After the 30-minute, 70°C-incubation step, add 4µL of A+ directly into the DNA ligase at the bottom of the
Ligase tube.
4. Using a new tip, add 4µL of K+ to the solution in the Ligase tube.
5. Using a new tip, add 3µL of 5x ligation buffer directly into the solution at the bottom of the Ligase tube.
Discard the buffer tube.
6. Add 2µL of dH2O to the Ligase tube, using a clean tip. Gently and slowly pump the plunger in and out to
mix the reagents. Do this without splashing the solution onto the sides of the microfuge tube. The table below
summarizes the contents of the Ligase tube.
7. If you have droplets of liquid clinging to the sides of the tube, briefly centrifuge the tube to pool the
reagents.
8. Place your ligase, A+ and K+ tubes in the microfuge racks at the front of the room. Your ligase tube will
be kept overnight at room temperature.
Conclusions
1a. Why was it important to place the A+ and K+ tubes in the 70°C water bath before setting up the ligation
reaction?
1b. What do you think might have happened if this step was omitted?
2. Make a diagram to show how the following sticky ends would join together.
(: = hydrogen bonding) See page 3.2 for base pairing example.
3. Although many recombinant plasmids are possible, draw three possible recombinant plasmids. Include as one
of the three the combination in which we are most interested—the one that combines pARA with the pKAN-R
fragment carrying
the rfp gene.
4. Could two rfp fragments join together and circularize in the Ligase tube?
5. In the DNA molecule, there are two kinds of chemical bonds: covalent chemical bonds and hydrogen bonds.
Briefly describe how these bonds differ in strength and where, in the DNA molecule, you would find them.
6a. During ligation, which of the bonds (hydrogen or covalent) form first?
Where do they form?
Which bonds form next and where do they form?
6b. DNA ligase is required to form which bond?
Lab4:
Confirmation of Restriction and Ligation
It is important, at this stage of our experimental procedure, that we confirm BamH I and Hind III have
digested the original pKAN-R and pARA plasmids and the restriction fragments have been ligated together by
DNA ligase. This lab will provide evidence that we have recombinant DNA molecules.
Gel electrophoresis is a procedure commonly used to separate fragments of DNA according to their molecular
size. Like the dyes you separated in Lab 1, DNA fragments will migrate through the agarose maze. DNA,
because of the phosphate groups, is negatively charged and will move toward the positive (red) electrode.
Because it is easier for small molecules to move through the agarose matrix, they will migrate faster than the
larger fragments. Picture a group of cross-country runners who are racing through a dense tropical rain forest.
All other factors being equal, the shorter runners will be able to navigate through the tangle of overhanging
vines and dense foliage faster than the taller runners. So, smaller DNA fragments will move through the tangle
of agarose molecules faster than the longer fragments.
We’ll take all of our plasmid samples: digested, undigested and ligated, and use electrophoresis to separate
these pieces. You might have predicted that your uncut plasmids would produce only a single DNA band;
there’s no reason why you would think otherwise. However, it is likely that two or three bands will appear in
the undigested plasmid lanes. The reason for this is that plasmids isolated from cells exist in several form s.
One form of plasmid is called “supercoiled.” You can visualize this form by thinking of a circular piece of
plastic tubing that is twisted. This twisting or supercoiling results in a very compact molecule, one that will
move through the gel very quickly for its size.
A second plasmid form is called a “nicked-circle” or “open-circle.” Often a plasmid will experience a
break in one of the covalent bonds located in its sugar-phosphate backbone along one of the two nucleotide
strands. Repeated freezing and thawing of the plasmid or other rough treatment can cause the break. When
this break occurs, the tension stored in the supercoiled plasmid is released as the twisted plasmid unwinds.
This circular plasmid form will not move through the agarose gel as easily as the supercoiled form. Although
it is the same size, in terms of base pairs, it will be located closer to the well that the supercoiled form.
The last plasmid form we are likely to see is called the “multimer.” When bacteria replicate plasmids, the
plasmids are often replicated so fast that they end up linked together like links in a chain. If two plasmids are
linked, the multimer will be twice as large as a single plasmid and will migrate very slowly through the gel. In
fact, it will move slower than the nicked-circle. Your pKAN-R – and pARA – samples, then, may each have
three bands that appear in the gel. Starting closest to the well, you might observe a multimer, followed by a
nicked-circle band and, finally, a fast traveling supercoiled band.
We will use a special staining technique that permits us to see the fragments embedded within the gel, then
make a photographic record of your gel to document this important step.
Materials
Reagent
Plasmid samples:
K-, K+
A-, A+
Ligated plasmid (“LIG” tube)
5x loading dye
1x SB (or 0.5x TBE)
DNA size marker (25 ng/µL)
0.8% agarose gel
Equipment & supplies
P-20 Micropipette and tips
1.5 mL microfuge tubes
Electrophoresis apparatus
Power supply
Marker pen
Plastic microfuge tube rack
Methods
1. Collect the five plasmid samples and the DNA marker from your teacher and place them in your plastic
tube rack. You should have six tubes.
2. Obtain five clean 1.5 mL microfuge tubes and label them as follows: A-, A+, K-, K+, and L. The
microfuge tube with the marker should already be labeled.
3. The following table summarizes plasmid sample preparation for electrophoresis.
See “Hints” before setting up these tubes.
Hints:
--For example, to the tube labeled “A-,” add 4µL of pARA-, 4µL of dH2O and 2µL of loading dye. The
loading dye should be located in your plastic microfuge tube rack next to the dH 2O tube.
-- If you study this table, you’ll see that you can add water to all five tubes, then add the loading dye to all of
the tubes without changing the tip. Then, dispense the plasmid sample into each tube, changing the tip
each time to avoid contamination.
-- Save the “LIG” tube that contains your ligated plasmid; there should be about 10µL remaining in this
tube.
Important: Return the “LIG” tube to the collection rack, at the front of the room, as you will need it for the
next lab.
-- Centrifuge all samples to pool the reagents at the bottom of each tube. Be certain that the tubes are placed
in a balanced configuration.
4. Prepare the gel and electrophoresis box to receive these plasmid samples.
--Be certain the gel wells are oriented closest to the negative (black) electrode.
--Pour the 1x SB buffer (or 0.5x TBE) over the gel until there are no visible “dimples” breaking the surface of
the buffer over the wells. It’s important that the gel be completely under the SB buffer. However, you don’t
want so much buffer in the box to allow the electrical current to run through the buffer and not the gel.
5. Take your plasmid samples and marker to the gel, along with your pipette and tips.
Continued on next page...
Continued from last page..
6. You will share this gel with another group.
Unless your teacher has you load your samples in a different pattern, load your samples in the order indicated
below. Follow the loading directions that begin with step seven. If you load your sample in a different order,
be certain to record it in your notebook for later reference.
7. Using a clean tip, set your P-20 micropipettor to 10 µL. Aspirate 10 µL of your
“DNA size marker” and slowly dispense it into the well.
--As you do this, slowly lower the pipette tip below the
surface of the buffer directly over, but not into, the well.
Putting the tip into the well can damage the wall of the
well or puncture the bottom of the well.
These are not good things to do.
--Use two hands to steady the pipette. Slowly dispense the sample by pushing to the first stop of the pipette.
Because of the loading dye, the sample will have a greater density than the electrophoresis buffer. This will
allow the sample to sink into the well.
-- Important: While holding the button on the first stop, slowly remove the pipette tip from the gel box. If
you’ve loaded your sample correctly, the well will be filled with a blue-colored solution.
8. Continue this procedure with the plasmid samples, following the order indicated on page 4.3. Change the
tip for each sample. If you choose to load your samples in a different order, be certain to record the sample
order in your notebook.
9. Close the gel box lid tightly over the electrophoresis chamber. Connect the electrical leads to the power
supply. Be certain that both leads are connected to the same channel (same side) with the negative (black) to
negative (black) and positive (red) to positive (red).
10. On the power supply, set the voltage to 130-135v.
11. After two or three minutes, look at your gel and be certain that the purple dye (bromophenol blue) is
moving toward to positive electrode. If it’s moving in the other direction—toward the negative (black)
electrode—check the electrical leads to see whether they are plugged into the power supply correc tly.
12. Be certain that you return your “LIG” tube to the front of the room. This tube should contain your
recombinant plasmids and will be used for the next lab.
13. Your teacher will explain what to do with your gels, so listen carefully. If your lab time is short, you may
not have sufficient time to complete the electrophoresis. The yellow dye will need to run just to the end of the
gel, about 40–50 minutes.
Conclusions
These questions are to be answered after you’ve had an opportunity to analyze you r gel photograph.
1. How did your actual gel results compare to your gel predictions?
2a. Are there any bands, appearing in your gel photo, that are not expected?
2b. What could explain the origin of these unexpected bands?
3a. Do you see evidence of the three plasmid forms in the uncut lanes?
3b. Is there evidence of more than one form of multimer?
4. Why are the ligated plasmids so close to the well?
5. Two of the 702 bp pKAN-R fragments, rfp gene fragments, may form a circularized fragment because each
end of the fragments terminates in BamH I and Hind III sticky ends. Is there evidence of a circularized 1404 bp
fragment in the ligated lane?
Lab4a:
Confirmation of pARA-R Restriction Digest
The purpose of this protocol is to examine the restriction fragments that result from the double digestion of
pARA-R by BamH I and Hind III (Lab 2a). Gel electrophoresis is a procedure commonly used to separate
fragments of DNA according to molecular size of the restriction fragments or number of base pairs. Like the
dyes you separated in Lab 1, DNA fragments will migrate through the agarose maze. Because of its phosphate
groups, DNA is negatively charged and will migrate towards the positive (red) electrode. Because it is easier
for small molecules to move through the agarose matrix, they will migrate faster than the larger fragments.
Picture a group of cross-country runners that are racing through a dense tropical rain forest. All other factors
being equal, shorter runners would be able to navigate through the tangle of overhanging vines and dense
foliage faster than taller runners. So, smaller DNA fragments will move through the tangle of agarose
molecules faster than the longer fragments.
We’ll take both plasmid samples- digested and undigested- and use electrophoresis to separate these
restriction fragments. You might have predicted that your uncut plasmids would produce only a single DNA
band, there’s no reason why you would think otherwise. However, it is likely that two or three bands will
appear in the undigested plasmid lane (control). Here is the reason for this: plasmids isolated from cells exist in
several forms. One form of plasmid is called “supercoiled.” You can visualize this form by thinking of a
circular piece of plastic tubing that is twisted. The twisting or supercoiling of the plasmid results in a very
compact molecule, and one that will move through the gel very quickly for its size.
A second plasmid form is called a “nicked-circle” or a “relaxed-circle.” Often a plasmid will experience a
break in one of the covalent bonds located in its sugar-phosphate backbone along one of the two nucleotide
strands. Repeated freezing and thawing of the plasmid or other rough treatment can cause the break. When this
break occurs, the tension stored in the supercoiled plasmid is released as the twisted plasmid unwinds. This
circular plasmid form will not move through the agarose gel as easily as the supercoiled form. Although it is the
same size, in terms of base pairs, it will be located closer to the well than the supercoiled form.
The last plasmid form you may observe is called the “multimer.” When bacteria replicate plasmids, the
plasmids are often replicated so fast that they end up in linked together like links in a chain. If two plasmids are
linked, the multimer will be twice as large as a single plasmid and will migrate very slowly through the gel. In
fact, it will move slower than the nicked-circle. The undigested plasmid, pARA-R, sample may have three
bands that appear in the gel. Starting closest to the well, you might observe a multimer, followed by a nickedcircle band and, finally, a fast traveling supercoiled band.
We will use a special staining technique that permits us to visualize the fragments embedded within the gel,
and then make a photographic record of your gel to document this important step.
Materials
Reagent
Plasmid samples:
A– and A+ (from Lab 2a)
0.8% agarose gel
5 x loading dye
1 x SB (or 0.5x TBE)
DNA size marker (25 ng/µL)
Equipment & supplies
P-20 Micropipette and tips
1.5 mL microfuge tubes
Electrophoresis apparatus
Power supply
Plastic microfuge tube rack
Marker pen
Methods
1. Collect both plasmid samples and the DNA marker from your teacher and place them in your plastic tube
rack. You should have three tubes.
2. Add 2 µL of loading dye to the A+ and A- tubes. Take care not to contaminate the plasmid samples. The
loading dye will increase the density of each sample so the DNA will sink into the gel well. The loading dye
also contains visible dyes so we can track the progress of our samples during electrophoresis. The DNA size
marker already contains loading dye. Without creating bubbles, gently pump the pipette several times to mix
the loading dye with the plasmid samples. Remember to use a new tip for each plasmid sample to avoid
contamination.
3. Prepare the gel and electrophoresis chamber to receive these plasmid samples.
-- Be certain the gel wells are oriented closest to the negative (black) electrode.
-- Pour the 1x SB (or 0.5x TBE) buffer into the electrophoresis box until there are no visible “dimples”
breaking the surface of the buffer over the wells in the gel. It’s important that the gel be completely
submerged below the buffer but you don’t want so much buffer in the box as the electricity runs only through
the buffer and not through the gel.
4. Take your plasmid samples and marker to the gel, along with your pipette and tips. You may share this gel
with another group.
5. Unless your teacher has you load your samples in a different pattern, load your samples in the order
indicated below. Follow the loading directions that begin with step six. If you load your sample in a
different order, be certain to record it in your notebook for later reference.
Continued on next page...
Continued from last page..
6. Using a clean tip and set your P-20 micropipette to 10 µL.
Aspirate 10 µL of your “DNA size marker” and slowly
dispense it into the well.
--As you do this, slowly lower the pipette tip below the surface of the buffer directly over, but not into, the
well. Putting the tip into the well can damage the wall of the well or puncture the bottom of the well.
--Use two hands to steady the pipette. Slowly dispense the sample by pushing to the first stop of the pipette.
Because of the loading dye, the sample will have a greater density than the SB buffer or TBE. This will allow
the sample to sink into the well.
-- Important: While holding the button on the first stop, slowly remove the pipette tip from the gel box. If
you’ve loaded your sample correctly, the well will be filled with a blue colored solution containing you
sample.
7. Using a clean pipette tip, load 12 µL of your A- sample into
the adjacent well.
8. Change the pipette tip and load 12 µL of your A+ sample
into the next well.
9. Close the gel box lid tightly over the electrophoresis chamber. Connect the electrical leads to the power
supply. Be certain that both leads are connected to the same channel (same side) with the nega tive (black) to
negative (black) and positive (red) to positive (red).
10. On the power supply, set the voltage to 130-135 v.
11. After two or three minutes, look at your gel and be certain that the purple dye (bromophenol blue) is
moving towards to positive electrode. If it’s moving in the other direction- towards the negative (black)
electrode, check the electrical leads to see if they are plugged into the power supply correctly.
12. Your teacher will explain what to do with your gels, so listen carefully. If your lab time is short, you
may not have sufficient time to complete the electrophoresis. The yellow dye will need to run just to the end
of the gel, about 30-40 minutes.
Conclusions
1. Besides using electrophoresis to separate DNA fragments, it can be used to estimate their actual size. For
example, we might be looking for a gene and we suspect it is of a certain size. Electrophoresis can be used to
locate fragments in that size range. In order to do this, we would need to run a gel with a mixture of DNA
fragments of known sizes. This mixture, called a “marker” or “ladder,” serves as a control or a standard to
which we can compare the positions of other DNA bands in the same gel.
In the diagram, below, the “marker” lane contains 10 DNA bands of known sizes. The fragment sizes are
given below. Using this information and the plasmid map of pARA-R, predict the positions of DNA bands
produced the A- and A+. You might want to review the different plasmid forms described on page 4a.1 and the
pARA-R plasmid map described on page 2a.3.
2. Compare your gel photo with your prediction. Do you see any unexpected DNA bands?
3. Relative to the DNA ladder, between what two bands is the rfp gene located?
Is this where you would have predicted it to be located?
4. In the A- lane, do you see evidence of different plasmid forms?
Which plasmid conformation shape migrates the fastest? Which is the slowest?
5. Does the A+ lane indicate complete digestion? Explain your answer.
6. Which DNA fragment contains the ampr gene?
What is the size of this DNA restriction fragment?
Lab5:
Transforming Escherichia coli with a Recombinant Plasmid
Thus far, you’ve produced ligated, recombinant plasmids. Hopefully, some of these DNA recombinants
will have the 702bp pKAN-R fragment, the rfp gene, ligated into the large pARA restriction fragment. This
plasmid is referred to as pARA-R. Now, we want to get these recombinant plasmids into bacterial cells so that
we can get the cells to express rfp gene and make the mutant fluorescent protein.
The process of taking up foreign pieces of DNA, like a plasmid, into a bacterial cell is called transformation.
Transformation is a process that occurs in nature, although it is probably somewhat rare. A British medical
officer, Frederick Griffith first studied the process, in 1928. Bacteria usually pass on extra chromosomal genetic
material, like plasmids, during conjugation (bacterial sex) rather than relying on luck. But taking up plasmids
can provide bacteria with certain genes that confer selective advantage, for example, antibiotic resistance.
Under experimental conditions, however, it is possible to prepare cells so that about one cell in a thousand will
take in a plasmid from the surrounding environment.
There are several factors that determine transformation efficiency. Two of these are related directly to the
plasmid used for transformation. The larger the plasmid, the less likely it will be taken up by the bacterium.
Remember, in order for the bacterium to take in foreign DNA, the plasmid must pass through bacteria’s plasma
membrane and cell wall.
Therefore, small plasmids are more likely to pass through the bacterium’s plasma membranes (E. coli has two)
and its cell wall than large plasmids.
Plasmids can assume different shapes. The supercoiled form is the easiest to get into the cell while the nickedcircle or the multimer, two or more plasmids linked together, are more difficult. The ligation tube, containing
the recombinant plasmids you prepared, does not contain any supercoiled plasmids. Supercoiling of a plasmid
requires an enzyme that is found in the bacterial cell; it was not included in your ligation tube. The recombinant
plasmids you prepared are primarily nicked-circled, but there is a wide variation in sizes.
In nature, transformation is a relatively rare event. To increase our chances of getting our recombinant
plasmids into bacterial cells, we will use “competent” cells. When cells are “competent,” it means that they are
ready to receive plasmids. For the most part, you don’t find competent cells in nature; instead, cells have to be
made competent in the laboratory. One common way this is done is by soaking the cells in calcium chloride.
Remember that DNA is negatively charged. Do you remember why? The plasma membranes surrounding the
bacterial cell also contain phosphate groups and are negatively charged. The problem of trying to get negatively
charged DNA past a negatively charged membrane is that like electrical charges tend to repel each other. When
cells are made competent, they are suspended in a solution of calcium chloride because calcium ions (positively
charged atoms of calcium, Ca++) help to neutralize the negative electrical charges of the plasma membrane and
the plasmid. With these repulsive charges neutralized by the calcium ions, the plasmid DNA has an easier time
passing by the plasma membrane of the bacterial cell. The cells have already been made competent for you and
your teacher will give you an aliquot. You will, however, need to do the next step.
Now that we have the negative charges on the DNA and the plasma membranes neutralized, we need to create a
bit of a pressure difference between the inside and the outside of the bacterial cell. This is done by first getting the
bacteria really cold and then quickly putting them into warm water. This is called “heat shock,” and it creates a
situation in which the pressure outside the cell is a tiny bit higher than inside the cell. This pressure gradient will
help to move the plasmid DNA from the outside to the inside of the bacterial cell. Following this brutal treatment,
we’ll need to feed our bacteria and let them recover for a few minutes before we spread them onto agar plates.
Once the cells have recovered, you’ll take samples of these cells and spread them on a series of sterile agar
plates. One of these plates will contain only bacterial food; the plate contains no antibiotic. This plate is marked
“LB.” A second plate contains LB and ampicillin; this plate is marked “amp.” The third plate contains LB,
ampicillin and a simple sugar called arabinose; this plate is marked “ara.”
Ampicillin is an antibiotic that prevents bacteria from fully forming its cell wall. Cells that are not ampicillin
resistant cannot grow in its presence; the new cells simply rupture or lyse. If a cell receives an ampicillinresistant gene, ampr, it will produce a protein that will chemically destroy ampicillin and, therefore, will be able
to grow with ampicillin in its environment.
Arabinose, a simple sugar, is needed by the bacterium to express the rfp gene. If a bacterium takes up
pARA-R, arabinose helps the enzyme RNA polymerase, needed to transcribe the rfp gene, to align itself
correctly on the plasmid. This relationship will be discussed in the next lab.
Although the E. coli strain that you are using in these labs is
relatively benign, it is important that you use proper techniques when handling them.
Materials
Reagent
LIG tube (recombinant plasmids)
100 µL of competent cells (LMG)
350 µL of LB broth (sterile)
Crushed ice (In a styrofoam cup)
Agar plates, sterile
1 LB, 1 LB/Amp, 1 Lb/amp/ara
Equipment & supplies
P-20 micropipette and tips
P-200 micropipette and tips
42° C water bath
1 pack cell spreaders (shared)
Plastic microfuge tube rack
1.5 mL microfuge tubes
Marking pen
Methods
In order to make this lab run smoothly, it’s important that you know which tasks have been assigned to each
group member before the beginning of the lab. One member should prepare the ice and get the competent cells,
another can retrieve your ligated plasmids and another can get the agar plates, LB broth and clean microfuge
tubes.
1. Pick up two clean microfuge tubes.
Label one “P+” and the other “P-.”
2. Pick up a Styrofoam cup with crushed
ice and place one tube containing 100 µL
of competent cells into the ice. It’s important
that the cells remain at 0°C. Also, place your P+ and P- tubes into the ice.
3. Pick up your ligated plasmids from the microfuge tube rack labeled “LIG tubes.” Your “ LIG” tube
should be labeled with your group number and class period.
4. Set the P-200 pipettor to 50 µL (set to “0-5-0”) and place a clean tip onto its barrel. Very carefully
resuspend the cells by gently pumping the cells in and out two times. Hold the tube by the upper rim to
avoid warming the cells with your fingers.
5. Aliquot 50 µL of the resuspended cells into the prechilled P+ and P - microfuge tubes. Immediately
return the aliquoted cells to the wet ice. Hold the tubes by the upper rim to avoid warming the cells
with your fingers.
6. Using the P-20 pipette, add 10 µL of your ligated plasmid to the tube labeled “P+.” Gently mix the
plasmid with the cell suspension by pumping the cell suspension two times. Immediately return the P+
tube into the ice. Do not add plasmid to the P– tube. The cells in this tube will serve as the “plasmid
control.”
7. Keep the cells in ice for 15 minutes.
8. While the cells are incubating in ice, obtain the following:
One each of these agar plates:
LB, LB/amp (LB + ampicillin)
and
LB/amp/ara (LB + amp + arabinose)
9. Label the bottoms (plate containing the agar) of all three plates with your group number and class period.
Write small and on the edge of the plate. Then divide the LB and amp plates down the middle using two lines.
Label one half of each plate “P+” and the other half with a “P-.” See the diagram below. Do not divide the ara
plate.
10. Following the 15-minute incubation in ice, carry the ice cup containing the cells to the 42°C water
bath. Take the tubes from the ice and hold them in the water b ath for 45 seconds. After the 45-second
heat shock, place them back into the wet ice immediately for at least one minute.
11. After one minute, use the P-200 pipette to add 150 µL (set to “1-5-0”) of LB broth to the P- tube.
Cap the tube and gently flick the lower portion of the tube two or three times to mix.
12. Use a new tip and transfer 150 µL of LB broth to the P+ tube. Close the cap and gently flick the
tube to mix.
13. Obtain one package of sterile cell spreaders from your teacher. Two groups will share this package.
14. You are now ready to spread your bacterial cells onto the sterile agar plates.
a. Using the P-200 pipette (set to “0-5-0”), gently pump the pipette two or three times to resuspend
the cells then aspirate 50 µL of cells from the P- tube. Open the lid from the LB plate like a
“clamshell.” Dispense these cells on the half of the plate marked “P-.” Close the lid.
b. Resuspend the cells by gently pumping the pipette then aspirate a second 50 µL aliquot for the
LB/amp plate. Remember, you want to deposit the P– cells on the half of the plate you labeled “P-.”
Cover the plate.
c. Open the package of sterile cell spreaders at the end closest to the spreader handles. You will
share this package with another group. Remove only one spreade r, keeping the others sterile. Hold
the spreader by the handle and do not allow the bent end to touch any surface, as this will
contaminate the spreader. Close the package to avoid contaminating any of the other spreaders.
d. Open the lid to the LB plate, like a clamshell, and gently using a light, gliding motion
spread the cells across the surface of the agar, keeping the cells on the P – side of the
plate. Try to spread them evenly and along the sides of the plate as well.
e. Carefully spread the P- cells on the LB/amp plate using the same spreader and technique.
f. Place the used spreader into the biohazard bag.
g. Repeat steps 14 a-e to inoculate the LB and LB/amp plates with the P+ culture. Be certain to use
the “+” pipette and a new spreader to avoid contamination.
15. Now you’re ready to inoculate the LB/amp/ara plate.
a. Using the P-200 pipette (set to “1-0-0”), transfer 100 µL of the P+ culture onto the surface of the
LB/amp/ara plate. Deposit the 100µL of cells on several areas across the ag ar surface rather than a
single spot.
b. Lift the lid, clamshell style, and spread the cells evenly over the surface of the plate.
c. Gently rotate the plate beneath the P+ spreader so that the cells can be spread over the entire
surface of this plate. Try to get the cells spread along the wall of the plate as well.
d. Cover the plate when finished.
16. Allow all three plates to sit right-side up for five minutes.
17. Using colored tape, tape all three plates together and place them in the incubator, gel-side up. Be
certain that you have clearly labeled your plates with your group number and class period. You can
mark the tape to help you find them for the next lab.
18. Discard cell-contaminated waste: spreaders, cell tubes, pipette tips, by placing th em into the cellcontaminated waste bag provided by your teacher.
Conclusions
Answer questions 1-3 before seeing the results of your transformation.
1. Predict the growth, if any, on the following plates. Remember the cells from the P+ culture were given
recombinant plasmids while those from the P– were not. Use a “+” if you expect growth and a “-“ if you expect
no growth.
2a. What do all of the cells growing on the LB/amp and LB/amp/ara plates have in common?
2b. What single restriction fragment must they all contain to grow on plates with ampicillin?
3a. Would you expect that all of the cells growing on the LB/amp/ara plate were transformed with the same
plasmid? Explain.
3b. How might you determine which of the cells on the LB/amp/ara plate contain pARA-R, the recombinant
plasmid that you’ve made by ligating the rfp gene with the large pARA restriction fragment?
Answer these questions after viewing the results of your transformation.
4a. Use the following table to compare how your actual transformation results differed from your predicted
results. See page 5.5 for “predicted” results.
4b. If your actual results differed from your expected, propose some reasons that might explain these
differences.
5a. How many red colonies were present on your LB/amp/ara plate.
5b. Why did the red colonies only appear on this plate and not the LB/amp plate?
5c. Would you expect that some of the bacteria on the LB/amp plate were transformed with pARA-R? Briefly
describe how you might test your answer.
Lab5a:
Transforming Escherichia coli with pARA-R
The process of taking up foreign pieces of DNA, like a plasmid, into a bacterial cell is called transformation.
Transformation is a process that occurs in nature, although it is probably somewhat rare. A British medical
officer, Frederick Griffith first studied the process, in 1928. Bacteria usually pass on extra chromosomal genetic
material, like plasmids, during conjugation (bacterial sex) rather than relying on luck. But taking up plasmids can
provide bacteria with certain genes that confer selective advantage, for example, antibiotic resistance. Under
experimental conditions, however, it is possible to prepare cells so that about one cell in a thousand will take in a
plasmid from the surrounding environment.
There are several factors that determine transformation efficiency. Two of these are related directly to the
plasmid used for transformation. The larger the plasmid, the less likely it will be taken up by the bacterium.
Remember, in order for the bacterium to take in foreign DNA, the plasmid must pass through bacteria’s plasma
membrane and cell wall. Therefore, small plasmids are more likely to pass through the bacterium’s plasma
membranes (E. coli has two) and its cell wall than large plasmids.
Plasmids can assume different shapes. The supercoiled form is the easiest to get into the cell while the nickedcircle or the multimer, two or more plasmids linked together, are more difficult.
In nature, transformation is a relatively rare event. To increase our chances of getting our recombinant
plasmids into bacterial cells, we will use “competent” cells. When cells are “competent,” it means that they are
ready to receive plasmids. For the most part, you don’t find competent cells in nature; instead, cells have to be
made competent in the laboratory. One common way this is done is by soaking the cells in calcium chloride.
Remember that DNA is negatively charged. Do you remember why? The plasma membranes surrounding the
bacterial cell also contain phosphate groups and are negatively charged. The problem of trying to get negatively
charged DNA past a negatively charged membrane is that like electrical charges tend to repel each other. When
cells are made competent, they are suspended in a solution of calcium chloride because calcium ions (positively
charged atoms of calcium, Ca++) help to neutralize the negative electrical charges of the plasma membrane and
the plasmid. With these repulsive charges neutralized by the calcium ions, the plasmid DNA has an easier time
passing by the plasma membrane of the bacterial cell.
Now that we have the negative charges on the DNA and the plasma membranes neutralized, we need to create
a bit of a pressure difference between the inside and the outside of the bacterial cell. This is done by first getting
the bacteria really cold and then quickly putting them into warm water. This is called “heat shock,” and it
creates a situation in which the pressure outside the cell is a tiny bit higher than inside the cell. This pressure
gradient will help to move the plasmid DNA from the outside to the inside of the bacterial cell.
Once the cells have recovered, you’ll take samples of these cells and spread them on a series of sterile agar
plates. One of these plates will contain only bacterial food. The plate contains no antibiotic. This plate is
marked “LB.” A second plate contains LB and ampicillin; this plate is marked “amp.” The third plate contains
LB, ampicillin and a simple sugar called arabinose; this plate is marked “ara.”
Ampicillin is an antibiotic that prevents bacteria from fully forming its cell wall. Cells that are not ampicillin
resistant cannot grow in its presence, the new cells simply rupture or lyse. If a cell receives an ampicillinresistant gene, ampr, it will produce a protein that will chemically destroy ampicillin and, therefore, will be able
to grow with ampicillin in its environment.
Arabinose, a simple sugar, is needed by the bacterium to express the rfp gene. If a bacterium takes up pARA-R,
arabinose helps the enzyme RNA polymerase, needed to transcribe the rfp gene, to align itself correctly on the
plasmid. This relationship will be discussed in the next lab.
Although the E. coli strain that you are using in these labs is relatively benign, it’s important that you use proper
techniques when handling them.
Materials
Reagent
E. coli (LMG) plate
Crushed ice
10 µL pARA-R (10 ng/µL)
1 mL 50 mM CaCl2
1 LB plate
1 LB/amp plate
1 LB/amp/ara plate
Equipment & supplies
1.5 mL microfuge tubes
Sterile inoculating loop
Disposable cell spreaders (2)
Microfuge tube rack
Permanent marker
P-20 micropipette and tips
P-200 micropipette and tips
Beaker with disinfectant
42° C water bath
Methods
Preparing competent cells for transformation.
1. Bacterial transformation requires sterile techniques. It is essential that directions be followed precisely.
2. Use the marker to label one of the 1.5 mL sterile microfuge tubes P+ and the other tube mark P-. Plasmid
DNA will be added only to the P+ tube. The P- tube will represent a negative plasmid control.
3. Pick up a tube containing CaCl 2. Use your P-200 pipette and a clean tip to transfer 250 µL of CaCl 2 to the
P+ and P- tubes. Hint: Set the P-200 pipette to 125 µL and transfer two aliquots to each of the labeled tubes.
The pipette should read “1-2-5” in the display window.
4. Your instructor will provide the class with a Petri plate containing colonies of E. coli cells. Use a sterile
inoculating loop to gently scrape two or three large bacteria colonies from the Petri plate and transfer them to
the P+ tube. Knock the loop against the side of the tube to dislodge the colonies from the loop. Cap the tube
and vigorously drag the tube across the surface of your microfuge tube rack to suspend the cells in the CaCl 2.
Continue to do this until you can no longer see any visible clumps of bacteria. Place this tube into your
crushed ice.
5. Repeat this procedure using the same inoculating loop, but transfer the colony and suspend the cells in the
P- tube. Place this tube in the crushed ice. Place the inoculating loop in disinfectant or cell waste bag for
proper disposal.
6.Transfer 10 µL of plasmid (pARA-R) directly into the cell suspension in the P+ tube. Briefly finger
vortex the mixture by gently flicking the bottom of the microfuge tube with your index finger. Avoid
splashing the mixture on the sidewall of the transformation tube. Return the P+ tube to the crushed ice.
7. Incubate both tubes in crushed ice for 15 minutes. Be certain the tubes are in contact with the ic e. It is
important that the cells get very cold.
8. Obtain one each of the following plates: LB, LB/amp and LB/amp/ara.
9. Using a marker and straight edge, draw a line down the center of the LB and LB/amp plates, but not the
LB/amp/ara plate. Make this division on the bottom gel side of the two plates. Label a “P-” and a “P+” on
each half of LB and LB/amp plate bottoms and a “+” on the LB/amp/ara plate bottom.
Transforming E. coli with pARA-R
1. Following the 15-minute chilling in ice, heat shock the cells in both tubes using the following
procedures:
--Take the ice container containing the transformation tubes P+ and P - from step 7 to the 42° C water
bath. It’s important that the bacteria receive a distinctly abrupt change in temperature.
-- Hold both tubes into the water bath for exactly 45 seconds.
-- Return both tubes to the crushed ice immediately for at least one minute.
2. Following the one-minute cooldown, return both tubes to the test tube rack and maintain at room
temperature.
Spreading transformed cells on agar plates
1. Position the three plates in the following order: LB, LB/amp, LB/amp/ara.
2. Set the P-200 pipette to 50µL. The pipette will read “0-5-0.” Holding the P- (control) cells between your
thumb and index finger. Gently flick the bottom of the tube to resuspend the cells. Deposit 50 µL of these
cells onto the “-” half of the LB and LB/amp plates. Do not deposit these cells on the LB/amp/ara plate.
3. Open the package of sterile cell spreaders at the end closest to the spreader handles. You will share this
package with another group. Remove only one spreader, keeping the others sterile. Hold the spreader by the
handle and do not allow the bent end to touch any surface as this will contaminate the spreader. Close the
package to avoid contaminating any of the other spreaders.
4. Open the lid to the LB plate, like a clamshell, and gently using a light, gliding motion spread the cells
across the surface of the agar keeping the cells on the “–” side only
of the plate. Try to spread them evenly and along the sides of the plate as well.
Using the same spreader, repeat this spreading procedure for the LB/amp plate.
5. Discard the cell spreader in disinfectant or the cell waste bag following its use on the LB/amp plate.
6. After resuspending the cells in the “P+” tube, use a new tip and deposit 50 µL of cells to the “+” side of
the LB and LB/amp plates; deposit 150 µL of cells to the LB/amp/ara plate.
7. Using a new cell spreader, repeat steps 4 and 5, spreading the cells on the “+” side only of the LB and
LB/amp plates and over the entire surface of the LB/amp/ara plate.
8. Discard this cell spreader in disinfectant or the cell waste bag.
9. Use masking tape to keep your plates together. Place your name on the tape so that you can locate your
plates later. Place the plates upside down into a 37°C incubator.
10. Incubate the plates for 24–36 hours at 37°C.
Conclusions
Answer questions 1-3 before seeing the results of your transformation.
1. Predict the growth, if any, on the following plates. Remember the cells from the P+ culture were given the
plasmid while those from the P– were not. Use a “+” if you expect growth and a “–” if you expect no growth.
2a.What do all the cells growing on the LB/amp and LB/amp/ara plates have in common?
2b. What fragment of the pARA-R plasmid allows these cells to grow on these plates?
2c.What is the size, in base pairs, of this fragment?
3. On which plate(s) would you expect the cells to express the rfp gene?
Answer these questions after viewing the results of your transformation.
4a. Use the following table to compare how your actual transformation results differed from your predicted
results.
4b. If your actual results differed from your expected, propose some reasons that might explain these
differences.
5a. How many “red” colonies were present on your LB/amp/ara plate?
5b. Why did the red colonies only appear on this plate and not the LB/amp plate?
Lab6:
Preparing an Overnight Culture of Escherichia coli
The purpose of this lab is to start a bacterial culture that will produce a sufficient quantity of mutant
fluorescent protein to enable you to isolate and purify the protein. From your LB/amp/ara plate, your teacher
will select one of the red colonies transformed with the pARA-R plasmid and use it to inoculate an overnight
culture.
The gene for mFP was originally isolated from a sea anemone. The mFP is used extensively in research as the
protein can be fused to other proteins and then followed through the cell using fluorescent microscopy. The
original fluorescent protein gene was mutated to produce a molecule that fluoresces many times brighter. The
plasmid pARA-R was then engineered for gene expression.
The diagram below depicts the region of pARA-R containing the major control elements required to express the
rfp gene. It’s important for you to note that only a small portion of the pARA-R plasmid is represented in this
diagram and the DNA is depicted as a straight line rather than a circle. The diagram identifies three important
regions: 1) araC gene; 2) the promoter region (PBAD); and 3) the location of the rfp gene relative to the other
control elements in the plasmid.
The araC gene codes for a regulatory protein known as the “AraC protein.” The AraC protein is involved
with turning the rfp gene off and on. The above diagram summarizes the relationship between the AraC gene,
transcription, messenger RNA, translation and the araC protein.
The promoter site is that portion of DNA where regulation of rfp expression occurs. When there is no
arabinose in the bacterium’s environment, the AraC protein will physically bind to two regions of plasmid, the
promoter site and a region near the araC gene. This causes the DNA molecule to bend around, forming a loop.
When the DNA is in this configuration, mRNA transcription cannot occur as it prevents RNA polymerase from
binding to the promoter site. Without mRNA, the bacterium cannot produce mFP.
When arabinose is present in the bacterium’s environment, arabinose binds with the AraC protein, forming a
complex. This prevents the DNA loop from forming. The binding of arabinose also causes a change in the
protein’s conformation (shape) resulting in the formation of a small pocket that will help a third molecule, RNA
polymerase, to join the complex.
This complex of three molecules binds to the promoter site, and RNA polymerase is aligned on the DNA
molecule in a way that it can transcribe the rfp gene. This transcription produces mRNA, which is translated into
mFP. The AraC protein, then, serves a duel function: It can inhibit mFP synthesis by looping the DNA and
preventing RNA polymerase from binding to the promoter region, and it can turn on the rfp gene transcription
and, therefore, mFP production, if it binds to arabinose.
When the bacterium expresses the rfp gene and produces mutant fluorescent protein, the cell takes these mFP
molecules and concentrates them into inclusion bodies. Inclusion bodies are concentrated granules of mFP
molecules and are not bound by a membrane.
Materials
Reagent
2 mL LB/amp/ara culture of E. coli (lab 6)
Lysozyme (10 mg/mL)
Binding buffer, 4 M (NH4)2SO4
Column equilibration buffer, 2 M (NH4)2SO4
Column wash buffer, 1.3 M (NH4)2SO4
Elution buffer, 10 mM TE
20% Ethanol
10% Bleach or other disinfectant
TE (same as elution buffer)
Equipment & supplies
Centrifuge
P-200 pipette and tips
P-1000 pipette and tips
Chromatography column
Microfuge tube rack
1.5 mL microfuge tubes
Permanent marker
6 mL waste collection tube
Cell-contaminated waste bag
Methods
Preparation of cell lysate from the overnight liquid culture
1. Obtain 1 mL LB/amp/ara culture from your teacher.
2. Examine this culture. What color is the culture?
3. Place this tube into the centrifuge.
Important: You or your teacher will need to make certain the tubes have been placed in the rotor in a
balanced configuration before the centrifuge is turned on. Centrifuge the microfuge tubes for 5 minutes.
4. After the rotor has stopped, carefully remove your tube to avoid disturbing the cell pellet.
5. Determine the location of the mFP. Is it in the bacterial cell pellet, or in the supernatant (the liquid above
the cell pellet)?
6. Once you’ve determined the location of the mFP, carefully decant (pour off) the supernatant into the
beaker containing disinfectant. Do this without disturbing the cell pellet.
7. Obtain a second 1 mL aliquot of the overnight culture and repeat steps 3–6.
8. Pick up a tube of “Elution buffer” and “Lysozyme” from your teacher.
9. Invert the microfuge tube containing the cell pellet and, using a small piece of paper towel, try to wick
away as much of the liquid as you can from your microfuge tube without touching the cell pellet. Discard
the used towel in the “cell-contaminated waste” bag.
10. Using the P-1000 pipette (set at “0-2-5”) and a clean tip, transfer 250µL of elution buffer to the cell pellet.
Close the cap tightly.
11. Resuspend the cells by dragging the tightly capped microfuge tube briskly across the surface of the
microfuge tube rack. You may need to do this several times to resuspend the cells. Examine the tube carefully
to make certain there are no visible clumps of cells.
12. Using the P-200 pipette (set at “0-4-0”), transfer 40µL of lysozyme to the resuspended cells. Lysozyme is
an enzyme that digests the bacterial cell wall. This enzymatic digestion of the cell wall greatly weakens the
cell, and the cells will begin to lyse (break open). Finger vortex or drag the microfuge tube across the surface
of the tube rack to mix thoroughly. If time permits, incubate samples at
room temperature for 60 or more minutes before freezing.
13. Make certain that you have labeled this tube with your group number and class period. Give the tube to
your teacher. These cells will be placed into the freezer following the incubation.
Cells can remain frozen, at –20°C, until the next lab.
Purification of mutant flurescent protein from the cell lysate
Getting the materials
1. Organize your group for multi-tasking.
-- Person A checks to see if the following reagents are at your workstation. These reagents will be shared
with another group.
Binding buffer
Equilibration buffer
Wash buffer
20% ethanol (if your group is the last to use the column)
-- Person B collects the lysed cells from your teacher; these cells were frozen overnight. This person should
take the cells to the centrifuge to pellet the cell debris.
-- Person C collects the following supplies:
2• 1.5 mL microfuge tubes. Label one tube “mFP” and
the other “super”
1• 6 mL waste collection tube (This may be already in the plastic tube rack.)
Preparing the column
2. Set up your chromatography column as directed by your teacher, being careful not to dislodge the stopcock
attached to the lower portion of this tube.
3. Set the waste collection tube into the plastic microfuge tube rack. Carefully open the column by turning the
stopcock valve and allow the fluid to begin draining from the column. Leave about 1 mm of this liquid
above the resin bead to avoid drying out the column.
4. Using the P-1000 pipette (set at “1-0-0”) and a clean tip, add 3000µL (=3ml) of equilibration buffer to the
chromatography column. Add the buffer slowly to the side of the column so that it does not disrupt the surface of
the resin bed. Let the buffer slowly dribble down the side of the column.
5. Allow the equilibration buffer to drain from the column into the waste collection tube, but leave about 1 or
2 millimeters of buffer above the resin bed to prevent it from drying.
6. Your chromatography column is now ready for the mFP sample. While you are waiting for the mFP
sample, be certain that the fluid is not draining from the column. If the waste collection tube is filled with
liquid, this is a good time to dump the liquid down the sink.
Preparing the mFP sample
7. Centrifuge the cell lysate for five minutes to pellet the cell debris. You or your teacher will need to check
the rotor to be certain it is balanced before closing the lid and spinning.
Balancing these tubes before centrifugation is very important.
8. After centrifugation, pick up your microfuge tube. Examine the microfuge tube. Where is the mFP:
supernatant or cell pellet?
9. Without disturbing the cell debris pellet, carefully remove 200µL of supernatant using the P-200
pipette (set at “2-0-0”) and a clean tip. Do this without transferring any cell debris. If you dislodge the
debris pellet, you will have to centrifuge the tube again. Dispense the 200µL of clean, debris -free
supernatant into a 1.5 mL microfuge tube labeled “super”(one of your group members should have labeled
this tube).
10. Replace the pipette tip on the P-200 and add 200µL of binding buffer to the supernatant you dispensed in
the tube labeled “super.” Mix the binding buffer with the supernatant by gently pumping the solutions in and
out using this pipette.
11. Using the p-1000 pipette (set at “0-4-0”) and a clean tip, add 400µL of this solution, mFP
supernatant/binding buffer, to the prepared column using the same pipette you used to mix the solutions. Do
this without disturbing the surface of the resin bed by dispensing the solution down the side of the column.
12. Without allowing the column to run dry, open the stopcock and allow the solution in the column to drain
into the waste collection tube. Leave about 1or 2 mm of buffer above the resin bed.
Continued on next page...
Continued from last page...
13. Examine the column and locate the mFP. Is the mFP spread throughout the resin bed or does it appear to
be restricted to a single band?
14. Using the P-1000 pipette (set at “1-0-0”), add 1000µL (=1ml) of wash buffer gently down the side of the
column. Without allowing the column to run dry, allow this buffer to drain from the column, leaving 1 or 2
mm of buffer above the resin bed.
15. Examine the column and locate the mFP. Has the location of the mFP changed in the resin bed? The wash
buffer will elute some of the less hydrophobic proteins off the column. The wash buffer’s salt concentration is
less than the binding buffer but not so low as to cause the mFP to release from the resin.
16. Using the P-1000 pipette (set at “1-0-0”) and a clean tip, add
2 x 1000µL (=2ml total) of elution buffer gently down the side of the column. As the mFP begins to drip
from the tip of the stopcock, collect the protein in the tube labeled “mFP.” Collect only the red eluate into this
tube. Cap the tube when you have collected all the mFP.
17. After all the mFP has been collected, add 2000µL (=2ml) of equilibration buffer to the column using the
P-1000 pipette and a clean tip. This will help prepare the column for the next class. If you are the last group
to use the columns, add 4000µL (=4ml) of 20% ethanol to the column. Allow about 2 mL of the ethanol to
drain from the column into the waste collection tube.
18. Cap the column tightly.
19. The solution in the waste collection tube can be discarded down the sink.
20. All microfuge tubes, except the one containing your mFP, should be discarded in the cell -contaminated
waste bag.
21. Compare your tube with mFP tubes from other groups. Is there a difference in intensity of color from
sample to sample?
Conclusions
1. What characteristic of mFP is used as the basis for separation by column chromatography?
2. Following centrifugation of the cell lysate, was the mFP localized in the supernatant or in the cell debris
pellet?
3. When would the hydrophilic proteins have been eluted from the column?
4. Does the eluate containing your mFP appear less bright or brighter than it did in the cell lysate following
centrifugation?
5. If there is a noticeable difference in the intensity of the red color, what might account for the difference?
6. How might the column be adjusted or modified to increase the purity of the mFP sample?
7. Although this laboratory involved the expression of a sea anemone gene, cite some examples of human
proteins that could potentially be expressed and purified using similar methods.
Lab7:
Purification of mFP from an Overnight Culture
When scientists at a therapeutics company, like Amgen, have successfully identified a promising
therapeutic protein, two objectives would be to locate and isolate the gene that encodes the protein. Once
isolated, the gene is inserted into a plasmid so that the gene can be cloned, as additional copies of the gene will
be needed for ongoing studies. The rfp gene was cloned in a plasmid called pKAN-R. pKAN-R is a cloning
vector, a plasmid that has been engineered to replicate in high numbers within the bacterial cell.
Later, cloned genes are inserted into plasmids that have been engineered specifically for protein expression in
bacteria or other suitable organism. Such plasmids are known as expression vectors. pARA-R is an expression
vector and carries the cloned rfp gene in a specific plasmid location, which allows the bacterial cell to produce
mutant fluorescent protein.
Transformed cells are allowed to express the protein in an overnight culture and then lysed (broken open) to
release the newly synthesized protein from the cell. The protein is isolated from the other cytoplasmic proteins,
purified and tested for activity.
You have already completed much of the work that parallels this drug discovery scenario. The bacterial cells
that have been growing in the LB/amp/ara broth have been expressing mFP and are now ready to be lysed (day
one of Lab 7) and the mFP purified (day two of Lab 7) using column chromatography.
Mutant fluorescent protein is a molecule that is about 238 amino acids in size. The native (as it exists in
Discosoma) protein is shaped like a cylinder with the fluorescent region, called the fluorophore, located in the
center of the cylinder.
In order to purify a molecule from other proteins present in the cell, one needs to look at how groups of
molecules differ from one another and how these differences can be used to effect separation.
One molecular attribute commonly used in purification is protein hydrophobicity. The term hydrophobicity is
related to the behavior of a molecule in water. If a molecule is hydrophobic, it fears water while hydrophilic
molecules love water. For example, oils, waxes and fats are hydrophobic; they do not dissolve in water. Table
sugar and table salt are hydrophilic, and they dissolve quickly in water.
It is not uncommon for large molecules, such as proteins, to have regions that are hydrophobic and other
regions that are hydrophilic. If these proteins are placed in water, the hydrophobic regions tend to “bend away”
from water while their hydrophilic regions try to bend toward the water. To a large extent, it’s the bending of
the protein’s amino acid chain that is responsible for its overall conformation or molecular shape, with
hydrophobic regions “hiding” in the interior of the molecule and water-loving regions on the outside.
It’s important for you to know that a bacterial cell contains many different kinds of proteins. The diagram
below is greatly simplified as it indicates only a few kinds. The problem, however, is how do you separate a
single protein, like mFP, from all of the others? A typical bacterium may contain more than a 1000 different
kinds of protein. The use of the recombinant expression vector, pARA-R, will make mFP isolation somewhat
easier: The E. coli cells you have cultured will have been made to produce a disproportionately high
concentration of mFP.
Protein purification can use hydrophobicity to separate and purify protein molecules. One common
purification procedure that uses differences in hydrophobicity to separate proteins is called column
chromatography. Column chromatography uses a plastic or glass cylinder into which a separating medium,
referred to as “resin,” is placed. The specific type of resin used will vary depending on what type of protein is
being purified. In this lab, we will be using a resin bed consisting of small hydrophobic beads. Mutant
fluorescent protein is highly hydrophobic and when mFP is placed into a solution of high salt concentration, the
mFP molecule is distorted in a way that will cause the hydrophobic regions of the molecule to adhere to the
hydrophobic resin in the chromatography column. The hydrophilic proteins made by the cell continue down the
column, through the resin without sticking to the resin bed and are flushed away.
Once the mFP is trapped in the resin bed, the column can be washed with a solution of lower salt
concentration to elute (wash out) moderately hydrophobic molecules from the column. This column wash buffer
will have a slightly lower salt concentration than the solution used to bind mFP to the resin but not so low as to
wash the mFP from the resin. Finally, we can use a solution of very low salt concentration to elute or release the
mFP from the resin beads. Under low salt concentration, the hydrophobic regions of the mFP molecule point
toward the interior of the molecule, thus releasing the mFP from the hydrophobic resin in the column.
Industrial protein purification is much more complex than this mFP purification protocol, but the principles
employed by industry are similar. The mFP sample that you obtain from this purification does contain other
proteins. The procedure, however, has removed many of the other proteins present in the bacterium’s
cytoplasm.
Materials
Reagent
0.5 mL of 10% Chelex solution
Master mixes I and II
Equipment & supplies
Boiling water bath
Microcentrifuge
Thermal cycler
Microfuge tube
Permanent marker
Sterile toothpicks
Methods
Getting your sample ready…
1. Obtain a Chelex tube. Note that this tube is identified with a number and letter. Record this number and
letter in your notebook. Only you will know this anonymous code.
2. To collect buccal epithelial cells, use a sterile toothpick or yellow pipette tip to gently scrape the inside of
both cheeks. This procedure should be noninvasive, so don’t draw blood.
3. Transfer the cells that you have removed from the toothpick to the Chelex tube. Vigorously twirl the
toothpick with the Chelex resin to knock off the cells from the toothpick.
This is important; you want to get as many cells off the toothpick and into the Chelex tube as possible.
4. Close the Chelex tube tightly. Take the tube to the boiling water bath or 100°C hot block. Boil or heat the cells
for 10 minutes. This heating will lyse the cells and help to destroy some of the nucleases, which degrade the
DNA
5. Use the high-speed centrifuge to spin down the Chelex and cell debris.
6. Using the P-20 pipette and a clean pipette tip, carefully remove
20 µL of supernatant and place it into a clean 1.5 mL microfuge tube. Avoid aspirating Chelex beads as this
will inhibit the downstream PCR procedure. Label this tube with your personal, anonymous code (number).
7. If your sample is not used immediately, leave this sample at the front of the room in the rack labeled
“Genomic DNA Samples.” These samples will be placed into the refrigerator overnight and returned to you
for the next lab.
Your genomic DNA sample can be kept in the refrigerator at 4°C
or freezer at _–20°C until you are ready to run the PCR reaction.
Lab8:
Genomic DNA Extraction from Buccal Epithelial Cells
The purpose of this lab is to collect a DNA sample from the cells that line the inside of your mouth and to
use this sample to explore one of the most powerful techniques in molecular biology—the Polymerase Chain
Reaction (PCR). Although PCR has many applications, it is commonly used to produce many copies of a
selected gene segment or locus of DNA. In criminal forensics, for example, PCR is used to amplify DNA
evidence from small samples that may have been left at a crime scene. A skilled technician can even obtain a
DNA sample left by the tongue on the back of a postage stamp used to send a letter. DNA samples obtained in
this manner have been used for PCR in several high-profile criminal cases.
To obtain your DNA sample, you’ll use a toothpick to obtain some buccal epithelial cells. The cells will be
transferred to a solution containing Chelex beads. The Chelex beads will bind divalent magnesium ions
(Mg++). These ions often serve as cofactors for nucleases that will degrade your DNA sample and may
interfere with the enzyme (Taq polymerase) used in the reaction. By removing magnesium ions, the degradation
of genomic DNA by nucleases is reduced. This mixture will be placed into boiling water to lyse the cells and
liberate the DNA.
The mixture of your genomic DNA, cell debris and Chelex beads is then centrifuged to pellet the cell debris
and Chelex, while keeping your genomic DNA in the supernatant. This is a quick and easy way to separate
genomic DNA from the cell debris. The DNA sample, however, is far from pure as it contains proteins and
nucleic acids from organisms that were in your mouth at the time of sampling (mostly bacteria and food).
Generally, these contaminants do not inhibit PCR because the process uses specific primers, short segments
of DNA about 25 nucleotides in length that can be made to target only human genomic DNA. Therefore, if
the supernatant carries some foreign DNA, it should not interfere with the targeting of the human -specific
primers. A more detailed description of PCR and the role that primers play will be discussed later in this lab.
Although we are using buccal epithelia as a DNA source, other tissues could have been used. Here are some
DNA yields from other human tissues: Blood yields 40 µg/ mL; hair root yields around 250 ng/mL; muscle
yields around 3 µg/ mL; and sperm yields 3.3 pg/cell.
The second part of this lab involves the actual PCR. You will use the sample of genomic DNA you just
collected as a target for the PCR reaction.
Methods
1. Obtain the genomic DNA sample with your number and the PCR tube labeled with your anonymous code
number.
2. The PCR tube already contains Master mix I. Master mix I contains the two
primers that target the tPA locus, dNTP’s (deoxynucleotide triphosphates: ATP, TTP, CTP
and GTP), PCR buffer, molecular grade water (very pure) and Taq polymerase.
3. Using a clean pipette tip, add 5 µL of your genomic DNA to this PCR tube. Carefully add your DNA
sample directly into the 10µL of Master mix I. Do this without creating bubbles.
4. Carefully cap the PCR tube. This is a very thin walled tube so avoid crushing it, but make sure that the cap
is firmly seated over the opening of the tube.
5. Place your PCR tube into the ice bucket by the thermal cycler.
6. The instructor will add 10µL of Master mix II, containing MgCl2, just before placing your samples into
the thermal cycler. Taq polymerase, an enzyme from the bacterium Thermus aquaticus, requires Mg++ ions
as cofactors to activate it.
7. Discard your genomic DNA sample.
8. Your instructor will run the PCR reaction at another time.
9. After the PCR run, 15 µL of your PCR product will be loaded into a 2% agarose gel. The gel will be
stained and photo-documented. These steps will be done by your instructor.
Conclusions
These questions should be answered after you have seen the PCR results.
1. What is your genotype with respect to the tPA gene?
2. With respect to the tPA gene, how many genotypes are possible?
3. Complete the following table using the class information.
Begin by determining the number of each genotype present in your class.
4. Calculate the frequency (percentage) of each genotype present.
5. Calculate the frequency of each allele present in your class.
6. Compare your results with other classes.
Is there a difference in relative frequencies of genotypes and alleles?
GLOSSARY
Adenine A nitrogen-containing base, one member
of the base pair AT (adenine-thymine). Adenine is found in both DNA and RNA.
Agarose A highly purified polysaccharide (complex
carbohydrate) commonly used in gel electrophoresis.
See also: electrophoresis.
Aliquot A sample of some part of a whole
(e.g., a sample of plasmid or a sample of cells).
Allele Alternative form of a genetic locus; a single
allele for each locus is inherited from each parent (e.g., at a locus for flower color the result might be for purple or white
petals).
See also: locus.
Alu element A 300 base pair piece of DNA that
has been randomly inserted throughout the genome. It no longer appears to have any function in the human genome. This
specific DNA fragment is called the “Alu” element because it carries a recognition sequence for the Alu restriction enzyme.
See also: genome, recognition sequence, restriction enzyme.
Amino acid Any of a class of 20+ molecules that are
combined to form proteins. The sequence of amino acids in a protein and hence protein function are determined by the
genetic code.
Ampicillin A commonly used antibiotic of the
penicillin family. Ampicillin prevents new cell wall material from linking properly in bacteria. This weakened cell wall will
prevent the growth of new bacteria.
Amplification An increase in the number of copies
of a specific DNA fragment.
See also: cloning, polymerase chain reaction.
Amp r Symbol used to designate the gene for
ampicillin resistance. This symbol is located in the pARA plasmid.
See also: pARA, plasmid.
Antibiotic A substance that kills or prevents the
growth of cells.
See also: ampicillin, kanamycin.
AraC protein A protein that is required for
expression of mutant fluorescent protein in cells transformed with pARA-R.
Aspirate To draw in or suck in.
Autosome A chromosome that occurs in
omologous pairs in both males and females and that does not bear the genes determining sex
ß -lactamase An enzyme encoded by the ampr gene
that destroys ampicillin.
Base One of the nitrogen-containing molecules that
distinguish one nucleotide from another. In DNA, the bases are adenine, guanine, cytosine and thymine.
See also: nucleotide, base pair,
base sequence.
Base pair Two nitrogenous bases (adenine and
thymine or guanine and cytosine) held together by hydrogen bonds. Two strands of DNA are held together in the shape of a
double helix by the bonds between base pairs. The sum of the base pairs in a DNA molecule is frequently used to express the
size of the molecule.
Base sequence The order of nucleotide bases in
a DNA molecule; determines structure of proteins encoded by that DNA.
Biotechnology A set of biological techniques
developed through basic research and now applied to research and product development. In particular, biotechnology refers
to the use by industry of recombinant DNA, cell fusion and new bioprocessing techniques.
Buffer A solution used to maintain an optimal
physical chemical environment for a chemical reaction. Buffers are used in restriction digests, ligations and for PCR.
Cancer Diseases in which abnormal cells divide and
grow unchecked.
Carcinogen Something that causes cancer to occur by causing changes in a cell’s DNA.
Carrier An individual who possesses an
unexpressed, recessive allele.
Cell The basic unit of any living organism that
carries on the biochemical processes of life.
Cell wall A structure that provides cells with
physical support; surrounds bacterial cells.
Chimera A mythical animal made from several
different animals. In molecular biology, it often is used to describe a recombinant DNA molecule.
Chromatography column An instrument that is
used to separate a mixture of molecules.
Chromosome In eukaryotes, a linear strand
composed of DNA and protein, located in the nucleus of a cell, that contains the genes; in prokaryotes, a circular strand
composed solely of DNA. In humans, there are 23 pairs of chromosomes in body cells.
Cloning Using specialized DNA technology to
produce multiple, exact copies of a single gene or other segment of DNA. A second type of cloning exploits the natural
process of cell division to make many copies of an entire cell. The genetic make-up of these cloned cells, called a cell line, is
identical to the original cell. A third type of cloning produces complete, genetically identical animals or plants.
Also see: cloning vector.
Cloning vector DNA molecule originating from a
virus, a plasmid or the cell of a higher organism into which another DNA fragment of appropriate size can be integrated
without loss of the vector’s capacity for self-replication; vectors are engineered to introduce foreign DNA into host cells,
where the DNA can be reproduced in large quantities. In the Amgen Labs, pKAN is used to clone many copies of the rfp
gene.
See also: expression vector, plasmid.
Codon Group of three mRNA bases that encodes a
single amino acid.
See also: amino acid, mRNA.
Competent Cells capable of taking up plasmid
DNA.
See also: Transformation.
Complementary base pair Nitrogen-containing
bases that are found opposite each other in a double-stranded DNA molecule. Complementarity is the result of size (a large
base must be opposite a small base) and number of hydrogen bonds between the adjacent bases in the pair (A and T form two
hydrogen bonds, G and C form three). Adenine is complementary to thymine and guanine is complementary to cytosine.
Covalent chemical bond One of the forces that
holds atoms together in a molecule. It is considered to be a strong bond and forms from the sharing of electrons between two
atoms.
Cytosine One of the nitrogen-containing bases
found in DNA and RNA. It is complementary to guanine.
See also: base pair, nucleotide.
Degrade To lower in quality; to convert into a more
simple compound; to decompose.
Denaturation The melting of DNA at high
temperatures into single nucleotide strands; changing the three-dimensional shape of a
protein molecule.
Deoxyribose A type of five-carbon sugar found in
DNA.
See also: DNA.
Directed evolution An experimental procedure
using PCR that causes the production of many random mutations. These mutations are then screened or analyzed for their
properties.
DNA (deoxyribonucleic acid) The molecule that
encodes genetic information. DNA is a double-stranded molecule held together by hydrogen bonds between base pairs of
nucleotides. The material of heredity.
See also: hydrogen bonds, base pairs, nucleotide.
DNA ligase An enzyme that joins DNA strands
by forming covalent chemical bonds in the sugar-phosphate backbone.
See also: covalent chemical bond.
DNA polymerase An enzyme used to replicate DNA
molecules. PCR uses a DNA polymerase from the bacterium Thermus aquaticus and is called Taq polymerase.
See also: DNA replication, polymerase chain reaction.
DNA replication The use of existing DNA as a
template for the synthesis of new DNA strands.
Double helix Structure in which two strands of
DNA are twisted spirally around each other.
Electrophoresis Movement of charged molecules
toward an electrode of the opposite charge; used to separate nucleic acids and proteins. When used to separate DNA fragments,
electrophoresis will separate the fragments by size with smaller fragments moving faster than smaller fragments.
Eluate The solution that washes out (e.g., solutions
that drip from chromatography column)
See also: chromatography column.
Enzyme A protein that acts to speed-up chemical
reactions.
Esherichia coli Common bacterium used in
numerous molecular biology protocols. The strain of E. coli used in the Amgen protocols is relatively harmless and does not
grow well outside the laboratory environment.
Eukaryote An organism that shelters its genes inside
a nucleus and has several linear chromosomes.
Exon Segment of a gene that encodes regions of a
protein.
See also: intron.
Express To make a protein.
See also: expression vector
Expression vector A plasmid genetically
engineered specifically to express genes (e.g., pARA).
See also: Cloning vector, plasmid.
Fluorescence The production of light by a molecule
(e.g., red fluorescent protein will release red light when exposed to ultraviolet light). The protein used in the Amgen Labs can
been seen with ambient light.
Forensics Specializing in the application of scientific
knowledge to legal matters. DNA is frequently used as evidence in legal matters.
Gamete Mature male or female reproductive cell.
Gene The fundamental physical and functional unit
of heredity; an ordered sequence of nucleotides located in a locus that encodes a specific functional product.
See also: gene expression.
Gene expression The process by which a gene’s
coded information is converted into the structures present and operating in the cell. Expressed genes include those that are
transcribed into mRNA and then translated into protein.
See also: mRNA.
Genetic code The sequence of nucleotides, coded in
triplets (codons) along the mRNA, that determines the sequence of amino acids in protein synthesis.
See also: codons, mRNA.
Genetic engineering Altering the genetic material
of cells or organisms to enable them to make new substances or perform new functions.
Genetic polymorphism Difference in DNA
sequence among individuals, groups or populations (e.g., alleles for the tPA locus amplified by PCR, or height in pea plantstall and short).
Genetics The study of inheritance.
Genome All the genetic material in the chromosomes
of a particular organism; its size is generally expressed as its total number of base pairs.
Genotype The combination of alleles an individual
carries for a specific trait.
See also: allele, phenotype.
Green fluorescent protein A protein produced by
the marine jellyfish, Aequoria victoria; protein encoded by the gfp gene.
Guanine One of the nitrogen-containing bases found
in DNA and RNA. It is complementary to cytosine.
See also: base pair, nucleotide.
Haploid A single set of chromosomes present in the
gametes of plants and animals. Humans have a haploid number of 23.
See also: diploid.
Heterozygous The two different copies, or alleles,
of the same gene.
See also: allele, gene, homozygous.
Homologous chromosomes Chromosome
containing the same linear gene sequence as another, each derived from one parent. While the gene sequence is (generally)
the same, the precise alleles may differ between chromosomes.
See also: allele, chromosome, gene.
Homozygous Having two identical copies, or a
leles, of the same gene.
See also: allele, gene, heterozygous.
Hydrogen bond A weak force resulting from the
attraction of a positive hydrogen atom to negatively charged regions of other atoms. This is the force that holds the two
strands of nucleotides together in the DNA molecule. The GC base pairs form three H-bonds while AT pairs for two.
Hydrophilic Water loving; dissolves in water; polar.
Some examples are sugar and salt.
Hydrophobic Water fearing; does not dissolve in
water; non-polar. Some examples are oil, wax and mutant fluorescent protein.
Intron Segment of a gene that does not code for a
protein. Introns are transcribed into mRNA but are removed before being translated into a protein.
See also: exons, mRNA, transcription,
translation.
Kanamycin An antibiotic that kills non-resistant
cells by inhibiting proteins synthesis.
kan r Symbol for the kanamycin resistant gene found
in the plasmid pKAN; encodes an enzyme called phosphotransferase that inactivates kanamycin.
Kilobase (kb) Unit of length for DNA fragments
equal to 1000 nucleotid pairs.
Kilobase ladder A set of standard DNA fragments
with lengths differing by one kilobase; used as a size standard in electrophoresis.
See also: electrophoresis.
Ligase The enzyme required to covalently join two
fragment of DNA.
Ligation The reaction that chemically joins two
fragments of DNA resulting a recombinant DNA molecule.
Locus A place or location on a chromosome, it may
be a gene or just any site with variations which can be measured. (e.g., Alu+, Alu- in the tPA intron).
Lysis To break open.
Marker See: kilobase ladder.
Messenger RNA (mRNA) The nucleic acid molecule
that carries genetic information from the genes to the rest of the cell for protein synthesis.
Mutagen Agent that can cause mutations.
See also: carcinogen.
Nitrogen-containing base A molecular component
of DNA and RNA nucleotides. In DNA there are four nitrogen-containing bases: A (adenine), G (guanine), T (thymine), C
(cytosine).
Nuclease A family of enzymes that will degrade
nucleic acids.
See also: degrade, enzyme.
Nucleic acid A large molecule composed of nucle
tide subunits; a polymer of nucleotides.
See also: DNA, nucleotide, polymer, RNA.
Nucleotide A subunit or monomer, of DNA and
RNA. Each nucleotide consists of a nitrogen-containing base, a five-carbon sugar and a phosphate group.
Operon A cluster of genes transcribed together to
give a single molecule of mRNA. The arabinose operon is used in the pARA expression vector to transcribe the rfp
gene.
Phage A virus for which the natural host is a
bacterial cell.
Phenotype Characteristic due to the expression of
our genes; usually refers to visible properties but may refer to characteristics revealed by laboratory test.
Plasmid Circular molecule of DNA which replicates
independently of the host’s genomic DNA (e.g., pKAN, pARA).
Polymer A large molecule made of similar or
identical subunits linked together (e.g., DNA, RNA, proteins).
Polymerase chain reaction (PCR) A chemical
procedure used to amplify a DNA sequence by repeated cycles of replication and denaturation.
Polymorphism Difference in DNA sequence at a
particular genetic locus. The differences may or may not result in a different phenotype.
See also: locus, phenotype.
Primer Short, preexisting polynucleotide chain to
which new nucleotides can be added by DNA polymerase. Two different primers are used to target the tPA locus amplified
by PCR.
Prokaryote Cell or organism with a single
chromosome and no nuclear membrane; bacteria.
Promoter Region of DNA in front of a gene that
binds RNA polymerase and so promotes gene expression; in the ara operon, the region of pARA that binds the AraC proteinarabinose complex and RNA polymerase prior to rfp expression.
Protein A large polymer of amino acids. Examples
are enzymes, mutant fluorescent protein and some hormones.
Recognition sequence (recognition site) Specific
nucleotide base sequence recognized by a restriction enzyme. The enzyme will cut the DNA within this nucleotide sequence.
Recombinant DNA molecule A combination of
DNA molecules of different origin that are joined using recombinant DNA techniques.
See also: ligation.
Restriction enzyme An enzyme that binds and cuts
DNA at a specific base sequence (e.g., BamH I and Hind III).
See also: recognition sequence
Restriction fragment The piece of DNA that
results from the cutting of the DNA molecule with a restriction enzyme. Fragments are often separated on a gel using
electrophoresis.
Restriction map Diagram of DNA, like a plasmid,
showing the restriction sites for restriction enzymes.
Ribose Five-carbon sugar found in RNA nucleotides.
Ribosome Site within the cell that assembles amino
acids into protein molecules.
SB buffer Sodium borate solution used as an
electrophoresis buffer. It is a solution that conducts an electric current and maintains a relatively constant pH.
See also: TBE.
Somatic cells Cells making up the body which are
not sex cells or gametes. These cells are usually diploid having two sets of chromosomes.
See also: gametes.
Sticky ends Ends of a DNA molecule cut with
certain restriction enzymes. These ends have unpaired bases.
Supercoiling Higher level of twisting of DNA often
found in plasmids.
Taq polymerase A heat stable enzyme commonly
used in PCR; polymerase found in the bacterium Thermus aquaticus.
See also: polymerase chain reaction.
TBE (Tris-Boric acid-EDTA) A solution used for gel
electrophoresis and in the preparation of agarose gels. The solution helps conduct an electric current while maintaining a
constant pH.
Thymine One of the bases found in DNA; the base
that is complimentary to adenine.
Transcription A chemical process that converts a
DNA nucleotide sequence into an mRNA nucleotide sequence; process that uses RNA polymerase to convert a DNA
template into a mRNA strand.
Transformation A process that places foreign DNA,
like a plasmid, into a cell.
Translation A chemical process that converts an
mRNA nucleotide sequence into an amino acid sequence; site of this reaction is the ribosome.
Vector See: cloning vector, expression vector.
Wild type The original or naturally occurring
version of a gene or protein.