Fall, 2005
TABLE OF CONTENTS
Biological Sciences 210 – Genetics and Molecular
Biology
Laboratory Manual
Fall, 2005
Project Pages
Project 1: Screening for P-Transposable Elements
in a Wild-Type
Strain of Drosophila
melanogaster 1
- 25
Project 2: Isolation and Characterization of
Mutations
In Drosophila melanogaster 26 - 37
Project 1: Screening for P-Transposable Elements in a
Wild-Type Strain of Drosophila
melanogaster
______________________________________________________________________________________________
Project
Summary
The genomes
of many wild-type strains of the fruit fly, Drosophila
melanogaster, contain P-elements.
P-elements, like other transposable elements, are a major cause of
mutation and play an important role in evolution. P-elements are short
stretches of DNA that can move around in the genome. Just how prevalent are P-elements in wild
fruit fly populations? Will every strain of D.
melanogaster isolated in the wild have the transposons? In an attempt to address this question, the
goal of this project is to determine whether or not a wild-type strain of D. melanogaster, isolated on this
campus, contains P-elements in its genome.
You will perform this project over the course of 8 lab sessions. You will isolate genomic DNA from wild-type
fruit flies and cut the Drosophila
genome into many small fragments using restriction enzymes. You will do Southern blot hybridization
analysis to probe for our sequence of interest, the Drosophila P-element. To
prepare for hybridizing your Southern blot, you will generate large amounts of
the probe DNA by transforming bacterial
cells with a recombinant plasmid containing P-element sequences, isolating the plasmid from the cells, and
purifying the P-element sequences by using the Polymerase Chain Reaction (PCR)
and gel electrophoresis. You will then use your pure probe DNA in an
attempt to identify P-elements on your Southern blot.
Introduction
Drosophila P-elements
The fruit
fly, Drosophila melanogaster, is an
ideal model organism for use in genetic and molecular analyses (for more
information on Drosophila melanogaster,
see Project 2). The genomes of many
wild-type strains of Drosophila
melanogaster contain P-elements.
P-elements are short stretches of DNA (<=2.9kb) that are transposable
elements, or transposons (or “jumping genes”); that is, they can physically
excise from one chromosome and move to another, or they can move from point to
point within a chromosome. P-elements
vary in length but are all derived from the 2.9kb complete P-element sequence,
which encodes a transposase (the enzyme that cuts out and moves its own stretch
of DNA). The complete P-element is actually more than just the transposase
gene, and its derivatives are usually incomplete; but for simplicity’s sake, we
will henceforth refer to the P-element as our “gene of interest.” Transposons are common in nature and
relatively easy to detect in a known DNA sequence, since the transposase works
by recognition of a characteristic transposon feature: flanking inverted repeats. When a transposon moves, oftentimes only a
portion of the DNA sequence will “jump” to a new location. Some of the gene sequence is usually left
behind in the original position.
Therefore, while P-elements can vary in length from 0.5-2.9kb, all are
recognizable by the 31bp flanking inverted
repeats (the “transposon footprint”).
Incomplete excision and movement of P-elements also leads to high copy
number, that is, to the presence of many copies of some or all of the P-element
scattered throughout the Drosophila genome. Since the goal of our experiment is to detect
P-elements in a wild-type population of Drosophila
melanogaster, their high copy number
suits our purpose well.
Recombinant DNA Technology
The
relatively recent development of recombinant DNA technology has enabled
biological researchers to make great strides in our understanding of the
structures and functions of genes.
Before the development of recombinant DNA technology, the complex
genomes of eukaryotes were extremely difficult to analyze. Recombinant DNA technology enables
researchers to break large genomes into specific fragments, which can then be
inserted into the small genome or into a DNA molecule from a different species,
such as a bacterium, and analyzed with relative ease. The small genome or DNA molecule into which
the fragments are inserted is called a vector, and recombinant DNA molecules
can be made by inserting DNA fragments from almost any species into a
vector. Recombinant DNA molecules are
commonly introduced into bacterial "host" cells by the process of
transformation. The vectors used to
construct recombinant DNA molecules are usually capable of replication, so once
inside a bacterial cell, the recombinant DNA molecule will be replicated,
resulting in the amplification (replication of many copies) of a specific DNA
fragment. The insertion of a fragment of
DNA into a vector, and the subsequent replication of the recombinant DNA
molecule is often called "cloning".
The ability to produce many copies of a given DNA sequence has been extremely
helpful in the analysis of gene structure and function. The Human Genome Project and other genome
projects would be impossible without
recombinant DNA technology. In addition,
genes encoding medically- and industrially-important polypeptides can be
inserted into vectors, and maintained and amplified in host cells. Host cells capable of synthesizing the
polypeptide products of these recombinant genes provide a means of producing
large quantities of important molecules.
For example, insulin, which is necessary for the treatment of some types
of diabetes, is produced inexpensively and in large quantities by bacterial
cells that express the human insulin gene from a recombinant vector. Recombinant DNA technology has been made
possible by the discovery and development of a number of important "tools"
- some of which are discussed below.
Restriction Enzymes
Restriction
endonucleases (or restriction enzymes), discovered in the 1970s, are valuable
tools for characterizing and manipulating DNA molecules. Restriction endonucleases are enzymes that recognize
specific nucleotide sequences, often 4, 6, or 8 base-pairs long, and cut DNA at
these sequences only. Each restriction enzyme recognizes its own, specific
sequence of nucleotides, called a "restriction site". There are many different restriction enzymes
that recognize and cut at many different nucleotide sequences. Restriction enzymes are produced by bacteria
as a defense against invasion from foreign DNA - especially from
bacteriophages. A bacterium modifies its
own DNA to protect the DNA from restriction enzymes. Foreign DNA that finds its way into the
bacterial cell will be recognized and digested, or "restricted" by
the cell's restriction enzymes. Restriction
enzymes are purified from bacterial cells for use in molecular biology
experiments.
By cutting a given piece of DNA
with specific restriction enzymes, we can determine the locations of the
restriction sites for those enzymes on that DNA, and generate a
"restriction map" of the given DNA. We can identify the
different-sized fragments resulting from the digestion of a given piece of DNA
with a certain restriction enzyme by "electrophoresing" the
restriction-digested DNA on an agarose gel, as shown in figure 1. Briefly, the digested DNA (consisting of a
mixture of many copies of each these three fragments) is loaded into one of the
sample wells at one end of an agarose electrophoretic gel. A voltage is set up across the gel such that
the sample wells are closest to the negative electrode, and the far end of the
gel is closest to the positive electrode. DNA has a net negative charge, and
thus migrates (moves) through the gel toward the positive electrode (this
process is called electrophoresis). The gel is composed of a porous matrix that
acts like a molecular sieve. Smaller DNA fragments move through the gel faster
than do larger DNA fragments. After the electrophoresis of the digested DNA has
been carried out for a certain period of time, it is stopped, and the DNA in
the gel is stained to make it visible.
The DNA is often visible as discrete bands. Each band represents a collection of many DNA
fragments that are all of the same size and have thus migrated the same
distance in the gel. Restriction enzymes
also enable us to break large genomes into specific, small fragments that can
be inserted into vectors to make recombinant DNA molecules.
Generation of Recombinant DNA Molecules
The combining
of two different DNA molecules is facilitated by the fact that many restriction
enzymes leave short regions of single-stranded DNA at their sites of
cutting. This results in the generation
of cohesive, or "sticky" ends at the restriction sites. If two DNA molecules are cut with the same
restriction enzyme, they will have complementary sticky ends that can be joined
together (ligated) by base-pairing with the help of enzymes called DNA
ligases.
Figure 2 shows an
example of a case in which a circular vector DNA, and human genomic DNA are
both cut with the restriction enzyme, EcoR
I, which recognizes the restriction site
5'-GAATTC-3'
3'-CTTAAG-5'
and cuts
between G and A on both strands, leaving short overhangs of single-stranded DNA
on either side of the restriction site.
If this particular vector has only one EcoR I site, the digest results in a linear vector with two sticky
ends, which looks like (N stands for a nucleotide of any type):
5'-AATTCNNNNNNNNNNNNNNNNNNNNNNNNNNG-3'
3'-GNNNNNNNNNNNNNNNNNNNNNNNNNNCTTAA-5'
A sticky end from one DNA molecule can link with, or anneal with a sticky end from another DNA molecule by virtue of complementary pairing between nucleotide bases in the single-stranded overhangs that are generated at the restriction sites:
These bases can pair with these bases
5'-NNNNNNNNNNNNNNG-3' 5'-AATTCNNNNNNNNNNNN-3'
3'-NNNNNNNNNNNNNNCTTAA-5' 3'-GNNNNNNNNNNNN-5'
The digestion
of human genomic DNA with EcoR I
results in a very large number of linear DNA molecules with sticky ends that
are complementary to the sticky ends of the digested vector DNA. Any one of these human DNA fragments can be
combined with the vector DNA by base-pairing between complementary sticky
ends. The digested DNA molecules are put
into solution together, and the sticky ends meet as the DNA molecules move
randomly about in the solution and bump into one another. Treatment with a DNA ligase enzyme will
covalently join the DNA molecules that have base-paired, to form one circular,
recombinant DNA molecule. The fragment
of DNA that is ligated to the vector DNA is called an "insert". Recombinant molecules can also be made by
digesting vector DNA with a mixture of two different restriction enzymes, and
digesting the DNA to be cloned with the same two restriction enzymes. This process is called
"directional" cloning, since the insert DNA will be spliced into the
vector DNA in a specific orientation, as shown in figure 3.
Vectors
Several
different types of vector can be used in the generation of recombinant DNA
molecules. These vectors originate from
bacteriophages (viruses that infect bacterial cells), bacteria, and also from
eukaryotes, such as yeasts. In this lab,
we will be using a bacterial vector, and this discussion will be restricted to
phage and bacterial vectors. A good
discussion of eukaryotic vectors can be found in Chapter 10 of the textbook,
· Plasmids
Plasmids are
small, circular DNA molecules. Plasmids
can be introduced into competent bacterial cells by transformation. Inside the bacterial cell, a plasmid exists
and replicates independently from the much larger bacterial genome, as depicted
in figure 4. Plasmids can (and have
been) engineered to carry genes that confer on the cells containing them
resistance to specific antibiotics.
Plasmids can also carry genes encoding certain enzymes that can be used
to "mark" bacterial cells by assaying the cells for the presence of
those enzymes. Since plasmids replicate
in bacterial cells, they allow amplification of the inserted DNA molecule into
many copies. One disadvantage of plasmid
vectors is they that cannot contain large inserts. Most plasmid vectors can hold only inserts
smaller than 10 kilobases (kb) (1 kb = 1,000 base-pairs).
·
Bacteriophage l Vectors
Derivatives
of the bacteriophage l can be used
to clone larger fragments of DNA - fragments on the order of 15 to 20 kb. The linear Phage l genome can be made into a cloning vector
by removing much of its central portion, which can then be replaced with
foreign DNA fragments, resulting in recombinant molecules. The recombinant phage are then replicated in
host E. coli bacterial cells.
· Cosmid
Vectors
Cosmid
vectors are hybrids between plasmid and phage vectors. Cosmids can be used to clone insert fragments
of up to 45 kb in length. Cosmids can be
maintained in bacterial cells in the circular plasmid form, and they can be
purified from the cells by packaging into phage particles.
Southern Blot Hybridization Analysis
One way to
check for the presence of a specific sequence (a partial or complete gene, for
example) in a genomic DNA sample is to perform Southern blot hybridization
analysis (When using genomic DNA, the technique is called genomic Southern blot
hybridization analysis). Like finding a
needle in a haystack, this powerful technique detects specific
DNA sequences
in the total DNA of an organism.
Southern blots can also be used for restriction mapping of DNA
sequences. The first step in genomic
Southern blot hybridization analysis is to digest genomic DNA from the organism
of choice with restriction enzymes and run it out on a gel. The next step is blot the gel onto a
nitrocellulose membrane, as shown in figure
5 . When you then “hybridize” this
membrane to a chemically labeled DNA probe specific for the sequence of
interest, the probe should stick – not just anywhere on your membrane, but
specifically where there is DNA of the matching sequence. A special labeling substrate is used to produce
light when it comes in contact with the labeled probe on the membrane (this is
called chemiluminescence), the light reaction can be used to expose X-ray film. When the X-ray film is developed, dark bands
will appear corresponding to the locations of DNA sequences that were bound to
the labeled probe on your membrane. A labeled
DNA size marker run on the same gel will allow you to measure the sizes (in
base pairs) of your element restriction fragments.
Polymerase Chain
Reaction (PCR)
Polymerase Chain Reaction (PCR) is a method for making many copies of, or amplifying, a specific DNA sequences (such as a particular gene or region of the genome). PCR is performed using a pair of specific primers, which are themselves single-stranded sequences of DNA, usually about 20 bases long. A primer is designed to be complementary in sequence to a specific gene, genomic region, or other DNA segment, and a pair of primers that flanks the specific DNA sequence is used to amplify (make many copies of) that DNA segment, which is called the template. The primers prime replication of the template DNA by an enzyme called Taq DNA polymerase, or Taq. Taq DNA polymerase is isolated from a bacterium called Thermus aquaticus, which lives in the very hot water of geothermal vents, and all of whose enzymes are active at high temperatures. Taq DNA polymerase is stable at 94oC and optimally active at 72oC. A PCR reaction takes place in a microfuge tube (also called an “Eppendorf” tube). The reaction mixture usually contains template DNA, primer pairs, Taq, free deoxynucleotide triphosphates (dNTPs – A, C, T, and G) and the appropriate buffers and salts, including magnesium – a necessary cofactor for the Taq enzyme. The microfuge tube containing the reaction mixture is placed in a thermocycler (a machine for varying temperature through a preset number of cycles. In PCR, genomic DNA is heated to denature the double-stranded DNA molecules, making them single-stranded (This is the Denaturation step). The reaction mix is then cooled, allowing primers to anneal to complementary sequences on opposite strands of the template genomic DNA (by hydrogen bonding between complementary bases (A-T, G-C) flanking the DNA segment to be amplified (This is the Annealing step). The reaction is then brought to an intermediate temperature, and, using free deoxyribonucleotides added to the reaction mixture, Taq DNA polymerase extends these primers from their 3' ends toward each other, as shown in figure 6 (This is called the Extension, or Elongation step). This replicates the region between the two primers, and generates two double stranded DNA molecules from the original one. After this round of replication is completed, the reaction mixture is heated to denature the double stranded DNA molecules, and then the temperature is lowered to allow the primers to anneal again - this time with double the number of templates. The reaction is then brought to an intermediate temperature again, allowing extension. This process is repeated for a number of cycles (usually 20-30 cycles), resulting in the production of many copies of the template DNA sequence. These copies are called the PCR product(s), or “amplicons”. If all goes as planned, the number of amplicons should double every cycle. So, given one molecule of template DNA, how many copies of a given amplicon should we have at the end of 30 cycles?
Depending on the number of nucleotide base-pairs present between the two primers used in a PCR reaction, different-sized fragments of DNA (products) will be generated in the reaction. We can identify the different-sized fragments of DNA resulting from PCR reactions by electrophoresing the product(s) of that reaction on an agarose gel.
This
Laboratory Project
Transformation of bacterial cells with
recombinant molecules
We have a
very small amount of a recombinant plasmid called pπ25.1 (kindly provided
by W. Engels). pπ25.1 contains
P-element probe sequences. We will be
using E. coli bacterial cells to
produce the mass quantities of this plasmid that we need. To do this, the plasmid will be introduced into E. coli cells by the process
of transformation. The bacterial cells
to be transformed must first be treated in a special way to make them
"competent" to take up foreign DNA.
In the process of transformation, under special conditions, competent
bacterial cells take up foreign DNA molecules from the surrounding medium.
Selection of cells that contain plasmids
The plasmid
vector that is part of the recombinant plasmid containing the P-element probe
is called pBR322. A simple map of pBR322
will be provided. pBR322 contains a gene
that confers resistance to the antibiotic ampicillin upon the cells that carry
the plasmid. Cells that carry pBR322 can
therefore be distinguished and purified from cells without pBR322 by growing
the mixture of cells with and without the plasmid on medium that includes ampicillin. Cells with pBR322 will thrive on the
ampicillin-containing medium, while the antibiotic will prevent cells lacking
the plasmid from growing on the plates.
PBR322 also contains another gene that provides resisitance to the
antibiotic tetracycline. In addition,
pBR322 has single restriction sites for many commonly-used restriction enzymes
(each of these enzymes will thus cut pBR322 in only one distinct site,
linearizing the plasmid). Many of these
restriction sites are within the tetracycline-resistance gene. Insertion of foreign DNA fragments (such as Drosophila P-element sequences) into these restriction
sites will thus disrupt the the tetracycycline-ressistance gene, rendering it
incapable of providing resistance to tetracycline. Cells that carry recombinant plasmids with
P-element sequence inserts in the the tetracycycline-resistance gene will not
grow on plates that contain tetracycline.
In
this project, you will plate cells that you have attempted to transform with pπ25.1,
and with the control plasmid pBR322 on two different types of nutrient agar
plates. The first type of plates will
contain ampicillin. The second type of
plates will contain tetracycline.
What do you
expect to see on each of the different plates you’ll be setting up?
Amplification of P-element-specific
sequences from the recombinant plasmids
After you
have purified your pπ25.1 plasmid DNA, you will set up a PCR reaction to
amplify the P-element sequences specifically.
You will design the pair of primers for this PCR reaction after
examining the nucleotide base-pair sequence of the P-element (available on the
World Wide Web). After the PCR reaction
is finished, you will run the PCR products on an agarose electrophoretic
gel. You will look for a band on the
gel that contains P-element sequences,
and actually cut that band out of the gel with a razor blade, thus preparing a
pure sample of P-element DNA to use a “hybridization probe” (see below).
Southern blot for the Drosophila
P-element
To check for
the presence of P-elements in your genomic DNA sample, you will perform a
Southern blot. You will first isolate
genomic DNA from the local strain of Drosophila
melanogaster. You will then digest
your genomic fly DNA 
with restriction enzymes and run it out
on a gel. Under the appropriate
conditions, you will then blot the gel onto a nitrocellulose membrane. You will then perform a reaction to label
your P-element hybridization probe with a molecule known as digoxigenin (“DIG”). This molecule is a derivative of the plant
steroid digoxin, and is attached to nucleotide residues. Your DNA is labeled using DNA polymerase in
the presence of these DIG-modified nucleotides.
You will then hybridize your labeled probe to your blot.
Detecting the
hybridized label requires a few steps.
First you will expose your blot to an Anti- DIG antibody that will bind
to your hybridized probe. We use a
special anti-DIG antibody that is attached to an alkaline phosphatase
enzyme. This enzyme can be used to generate
light using a chemiluminescent substrate.
You will use this substrate on your blot and the light reaction will
occur wherever your hybridized probe is bound.
Outline of the Lab Project
Lab Session 1
1) Transform E. coli cells with plasmid containing
probe DNA and plate the cells.
Lab Session 2
1) Begin
isolating genomic DNA from the fruit fly, Drosophila
melanogaster.
2) Identify
transformant colonies on your plates from Lab Session 1 and calculate
transformation efficiencies.
Lab Session 3
1) Finish
isolating genomic DNA from the fruit fly, Drosophila
melanogaster.
2) Digest the
genomic DNA with restriction enzymes.
Lab Session 4
1) Run a gel
on the digested Drosophila genomic
DNA and set up a Southern blot.
Lab Session 5
1) Your instructor
will grow 2 ml cultures of selected colonies from your plates that you set up
during Session 1.
2) You will
isolate recombinant plasmid DNA containing the probe from these cultured cells.
Lab Session 6
1) Run a gel
on the PCR products and isolate the probe DNA fragment.
Lab Session 7
1) Purify PCR product and
label with DIG
2) Set up hybridization
reaction.
Lab Session 8
1) Hybridize blot to
antibody,
2) Develop with
chemiluminescence
3) Expose to film
Experimental Procedures (work in groups of 2)
Lab
Session 1
Part 1. Transform E. coli cells with plasmid containing
probe DNA and plate the cells.
During this
lab session, you will begin the process of preparing large amounts of the
P-element probe DNA. You will introduce
a recombinant plasmid called
pπ25.1, which contains P-element
probe sequences, into competent E. coli
cells, "transforming" the bacterial cells. You will then plate the cells on nutrient
agar plates that contain the antibiotic, ampicillin, which will kill
untransformed cells. You will also be
doing some control experiments. You will
transform E. coli cells with a pure
plasmid called pBR322, which is the vector plasmid of pπ25.1 (pBR322 =
pπ25.1 – P-element sequences). You
will plate your two sets of transformed cells on two different types of
plates. One type of plate will contain
ampicillin, while the other will contain a different antibiotic called tetracycline. PBR322 contains genes for both ampicillin
resisitance and tetracycline resistance, so cells transformed with pBR322
should grow on both types of plates.
After observing restriction maps of pπ25.1 and pBR322, make a
prediction about which plates cells transformed with this recombinant plasmid
will grow on.
Procedure
1) Obtain two
microfuge tubes of plasmid DNA, labeled pBR322, and pπ25.1. Into each of two empty microfuge tubes, pipet
90µl dilution fluid (sterile saline).
Pipet 10 µl of pBR322 plasmid DNA sample into one of the tubes with 90µl
dilution fluid to make a 10-1 dilution. Label this tube “pBR322 DNA Only
control”. Repeat for the pπ25.1
plasmid DNA sample, then set these two diluted (10-1 ) DNA samples
aside for a while.
2) Obtain a
microfuge tube containing competent E.
coli cells, and place it on ice.
3) Gently tap
the tube of competent cells with the tip of your finger to ensure that the
cells are well suspended.
4) Using a
pipetter, transfer 0.2 ml (200 µl) of competent cells into the tube containing
what is left of the undiluted pBR322 plasmid DNA (approx. 5ul). Cap the tube,
and tap it with the tip of your finger to mix the cells with the DNA. Put the
tube on ice for 20 minutes. The competent cells, which are suspended in CaCl2,
will begin to take-up the recombinant plasmid DNA.
5) Similarly,
using a sterile pipet tip, transfer 0.2 ml (200 µl) of competent cells into the
remaining tube of undiluted pπ25.1 plasmid DNA. Cap and tap as before, and
put the tube on ice for 20 minutes.
6) Similarly,
using a sterile pipet tip, transfer 0.2
ml (200 µl) of competent cells into a tube labeled “Cells Only”, which contains
only 5 µl of sterile saline. Cap and tap as before, and put the tube on ice for
20 minutes.
7) After the
20 minute ice treatment, place the three tubes in a 42oC water bath
for 1.5 minutes. This heat shock facilitates the uptake of DNA by the bacterial
cells.
8) Transfer
the tubes to ice for 1.5 minutes.
9) Using a
sterile pipet tip, add 0.8 ml (800 µl) of LB (L Broth, a nutrient-rich
bacterial growth medium) to each tube, including the two “DNA Only control” tubes.
10) Incubate
the tubes at 37oC for 20 minutes, then tap each tube to mix its
contents well. Place them back at 37oC for an additional 20 minutes.
This incubation period allows the cells to recover from the CaCl2
treatment, and to begin expressing the ampicillin resistance and/or the
tetracycline resistance genes on the plasmid. Mix the contents of each of the
tubes by tapping .
11) You will
now dilute your transformation mixtures prior to plating them on selective agar
medium. Transfer 0.1 ml (100 µl) of the
pBR322 transformation mixture into a tube containing 9.9 ml of dilution fluid. This is a 1:100 (or 10-2)
dilution. Also make a 10-3
dilution by transferring 1.0ml of the 10-2 dilution into a tube
containing 9.0 ml of dilution fluid
12) Repeat
step 10 for the pπ25.1 transformation mixture.
13) Using the
"spread plating technique" (which your lab instructor will explain
and demonstrate for you), plate 0.2 ml (200 µl) of each of the three dilutions
of the pBR322 transformation mixture (undiluted, 10-2, 10-3)
onto LBAmp plates and also onto LBTet plates.
14) Repeat
step 12 for the three dilutions of the pπ25.1 transformation mixture,
using LBAmp and LBTet plates.
15) Plate 0.1
ml (100 µl) of the “Cells Only” on both an LBAmp and an LBTet plate. Also, plate 0.2 ml (200 µl) of each of the
two “DNA Only control” (10-1 dilutions of plasmid DNA samples + 800
µl LB) onto separate LB plates (plate with LB but no antibiotic).
16) There
should be a total of 16 plates altogether.
Incubate these plates upside-down at 37oC. Tomorrow morning, your instructor will move
the plates to the refrigerator for storage until next week.
17) In
addition, your lab instructor will to plate several dilutions (try 3 different
dilutions: 10-4, 10-5, 10-6) of untransformed
E. coli cells on nutrient agar plates
lacking antibiotic. This will
enable you to determine the titer (concentration) of the original suspension of
cells, and thus to calculate transformation efficiencies that you achieve.
Lab
Session 2
1. Begin
isolating genomic DNA from the fruit fly, Drosophila
melanogaster.
2. Identify
transformant colonies on your plates from Lab Session 1 and calculate transformation
efficiencies.
During this
lab session, you will also begin the process of isolating genomic DNA from
fruit flies. You will anesthetize about
50 adult flies, then grind them up (homogenize them) in a special buffer that
keeps DNA stable. You will then treat
the resulting homogenate with a detergent to disrupt the cell membranes, and a
protease enzyme to destroy proteins.
Finally, you will mix the homogenate with a combination of phenol,
chloroform, and isoamyl alcohol, and store it in the freezer until next lab
session.
You will also
examine the plates that you prepared last session, looking for bacterial
colonies. You will count the numbers of colonies on each of your
plates and determine transformation efficiencies.
Part 1.
Isolation of Genomic DNA from Drosophila
melanogaster
Procedure
1) You will
first be provided with a vial containing wild-type or laboratory strains of fruit
flies. Anesthetize all of the flies in
the vial as demonstrated by your lab instructor. Place the anesthetized flies on a white paper
card, and view them using a dissecting microscope. For more information about fruit flies, see
Project 2.
2) Practice
moving the flies around on the white paper card with a fine paint brush. Transfer approximately 50 flies into a 1.7 ml
microfuge tube. Keep the tube on ice.
3) Add 500 µl
of HOM* buffer to the flies in the microfuge tube. Using a blue plastic homogenizer, homogenize
the flies. Use several strokes, and do
not be too rough. Add another 500 µl of
HOM buffer to the tube and continue to homogenize the flies until no body parts
are recognizable.
(*HOM: 80mM EDTA, 100mM Tris-Cl, 160 mM sucrose, pH
8.0)
4) Transfer the homogenate to a 15 ml centrifuge
tube. Add another 500 µl of HOM to the
original microfuge tube, then transfer this to the 15 ml centrifuge tube.
5) Add 75 µl
of 10% SDS and 25 µl of 10 mg/ml Proteinase K.
6) Incubate
this for at least 1 hour at 68oC.
During this incubation, do Part 2 of today’s lab session.
7) After the
68oC incubation, cool it on ice until it comes down to room
temperature.
8) Add 1
volume (~2 ml) of phenol:chloroform:isoamyl alcohol (25:24:1) and mix. Store this in the freezer until next lab
period.
Part 2.
Examination of Transformation Plates
1) Obtain the
plates that you prepared last session.
2) Your
instructor will show you how to effectively count the number of colonies on
each plate, and determine the transformation efficiencies.
Finish Part 1 of today’s lab session.
Lab
Session 3
1. Finish
isolating genomic DNA from the fruit fly, Drosophila
melanogaster.
2. Digest the
genomic DNA with restriction enzymes.
During this
lab session, you will complete the process of isolating genomic DNA from fruit
flies. You will perform a series of
phenol:chloroform:isoamyl alcohol extractions to remove impurities from the
DNA, and you will precipitate the DNA away from other impurities with
ethanol. You will dissolve your fly genomic DNA in a buffer
called TE. Finally, you will digest your Drosophila
genomic DNA with restriction endonucleases.
Procedure
1) Remove
your sample from lab session 2 from the freezer, and allow it to thaw at room
temperature.
2) Centrifuge
your sample at 5,000 rpm for 5 minutes.
3) Transfer
the aqueous (top) layer to a clean 15 ml centrifuge tube, and add 1 volume of
phenol:chloroform:isoamyl alcohol (25:24:1) to it. Mix, and centrifuge at 5,000 rpm for 5
minutes.
4) Transfer
the aqueous (top) layer to a clean centrifuge tube, and add 1 volume of
chloroform to it. Mix, and centrifuge at 5,000 rpm for 5 minutes.
5) Transfer
the aqueous (top) layer to a clean centrifuge tube, and add 1 volume of
chloroform to it. Mix, and centrifuge at 5,000 rpm for 5 minutes.
6) Transfer
the aqueous (top) layer to a clean centrifuge tube, and add 225 µl of 8M
potassium acetate (KOAc) to it. Mix, and place this on ice for at least 30
minutes.
7) Centrifuge
this at 13,000 rpm for 20 minutes.
8) Transfer
the supernatant to a clean tube, and add 1 volume of 95% ethanol to it. Incubate this for 10 minutes at room
temperature to precipitate the Drosophila
genomic DNA.
9) Centrifuge
this at 10,000 rpm for 10 minutes.
Remove the ethanol, and allow the pellet to air-dry slightly.
10) Resuspend
the pellet in 50 µl of TE buffer.
11) Transfer
all of the resuspended DNA into a clean microfuge tube and label the tube
appropriately.
12) Pipet 14 ml of the DNA into a microfuge tube containing restriction
digestion mix. Label the tube “Digested
DNA”.
Save the rest of your carefully labeled Drosophila genomic DNA in the
freezer.
13) Tap the
tube gently to mix the contents, and place it at 37oC until tomorrow
morning. Your lab instructor will remove
it and place in the freezer.
Lab
Session 4
1. Run a gel
on the digested Drosophila genomic
DNA and set up a Southern blot.
During this
lab session , you will run your restriction-digested Drosophila genomic DNA on an electrophoretic gel. You will then set up a Southern blot to
transfer the DNA on the gel to a nitrocellulose (or nylon) hybridization
membrane.
Part 1.
Electrophoresis
Procedure
1) Remove the
tube containing your restriction-digested Drosophila
genomic DNA (from Lab Session 3) from the freezer, and place it in a 70˚C
water bath for 5 minutes.
2) You will
be provided with another microfuge tube, labeled "l-Hind
III". This tube contains phage l DNA completely digested with Hind III. This digested DNA
will consist of DNA fragments of known length, which will serve as DNA
size-standards on your electrophoretic gel.
Place this tube in a 70˚C
water bath for 5 minutes, then keep it on ice until you are ready to use
it.
3) Add 5 µl
of electrophoresis sample buffer to the tube containing your restriction-digested Drosophila genomic DNA. Tap the tube to mix the contents.
Caution: The gel and reservoir buffer used in
steps 4-8 contain ethidium bromide (etBr), a carcinogen. Wear gloves when handling anything containing
etBr!
4) Using your
pipetting device, load the entire contents of the "l-Hind
III", and the tube containing your restriction-digested Drosophila genomic DNA into the sample
wells of an agarose gel as shown (2 pairs of students will share each gel. One pair of students will use wells 1 - 4. The other pair of students will use lanes 5 -
8:
Sample Well
Number
(Starting
at
left) Sample
1 "l-Hind
III"
2 Leave
Empty
One Pair of
Students 3 Digested Fly DNA
4 Leave
Empty
________________________________________________________________________
5 "l-Hind
III"
6 Leave
Empty
Second Pair
of Students 7 Digested
Fly DNA
8
Leave
Empty
5) Cover the
electrophoresis unit and plug the leads into the power supply. The leads are
color coded so that the red lead plugs into the positive terminal, and the
black lead plugs into the negative terminal. The sample wells in the gel should
be closest to the negative (black) electrode.
6) Turn the
power source on and set the voltage at about 120 volts. Electrophorese until
the bromphenol blue (dark blue dye) in the samples has migrated to within 5 mm
of the positive (red) electrode end of the gel. At 120 volts, this should take
about 1 hour.
7) After your
gel run is complete, turn the power supply off and unplug the leads. Remove the
gel from the unit and place it on a ultraviolet (UV) transilluminator.
8) Turn the
UV transilluminator on, and photograph your gel using a gel imager as demonstrated
by your instructor.
Part
2. Southern Blot
1) Rinse the
gel in distilled water.
2) Soak the
gel in Southern Denaturing Solution (0.5N NaOH, 1.5M NaCl) for 15 minutes on a
gentle shaker. Repeat using fresh
Southern Denaturing Solution for another 15 minutes.
3) Soak the
gel in Southern Neutralizing Solution (1M Tris-HCl, 1.5M NaCl, pH=8.0) for 15
minutes on a gentle shaker. Repeat using
fresh Southern Neutralizing Solution for another 15 minutes.
4) During
soaks, prepare a Southern blotting stack as follows:
a. Place a sponge in the center
of a Tupperware dish.
b.
You will be provided with a piece of Whatman 3MM (or equivalent) filter paper
cut to the exact size of the sponge. Place
this piece of filter paper on top of the sponge.
c.
Fill the dish with 20X SSC so as to completely soak the Whatman paper. This should lie flat against the sponge.
d.
You will be provided with a piece of nitrocellulose or nylon transfer membrane
cut to the exact size of the gel.
IMPORTANT: Never handle membrane without
gloves. Label the dry membrane with a
fine-point Sharpie.
e.
You will be provided with two pieces of Whatman paper cut to the exact size of
the gel. Place them and the membrane in
2X SSC.
5) When soaks
are finished, place the gel upside-down onto the filter paper that covers the
sponge. Remove any air bubbles from
beneath the gel, using a glass roller.
6) Lay the
wetted transfer membrane onto the gel, being careful to roll out any air
bubbles. With a razor blade, cut off one
corner – approximately 0.5cm2 – of both gel and membrane.
7) Lay the
two pieces of wetted Whatman paper onto the transfer membrane. Remove bubbles.
8) Place a
stack of paper towels approximately 1.5” thick on top of the filter paper.
9) Place a
glass plate and appropriate weight (ask instructor) on top of the paper
towels. Allow to transfer overnight.
Lab
Session 5
1. Your
instructor will have grown 2 ml cultures of selected colonies from your plates
that you set up during Session 1.
2. You will
isolate recombinant plasmid DNA containing the probe from these cultured cells.
3. Set-up a
PCR reaction to specifically amplify P-elment sequences from the recombinant
plasmid. Save these PCR products for
electrophoretic analysis during Session 6.
On the
afternoon prior to this lab session, your instructor selected bacterial colonies from your plates that you
set up during Session 1. These bacterial colonies consist of cells that were
transformed with the recombinant plasmid pπ25.1 (contains P-element probe
sequences). Your instructor inoculated 2
ml of LB (nutrient broth) + ampicillin with each colony. The inoculated cultures were grown overnight
at 37oC. During this lab
session , you will isolate plasmid DNA
from these overnight liquid cultures.
You will then set up a PCR reaction to amplify P-element sequences for
use as a hybridization probe for your Southern blot.
Part 1.
Isolation of recombinant plasmid DNA
Procedure
1) Each pair
of students should obtain two test tubes containing 2 ml overnight cultures,
each started from a single colony of cells transformed with the recombinant
plasmid pπ25.1 (contains P-element probe sequences).
2) Pour 1.5
ml of one overnight into an approporiately-labeled microfuge tube. Pour 1.5 ml of the other overnight culture
into a second appropriately-labeled
microfuge tube. The following
instructions apply to each of the two microfuge tubes.
3) Centrifuge
the tube at full speed for 20 seconds to pellet the cells.
4) Pour the
supernatant out of the tube. Discard the
supernatant.
5) Resuspend
the bacteria in 100 µl (0.1 ml) of GTE buffer by vortexing well.
6) Add 200 µl
(0.2 ml) of NS solution to each tube to lyse the bacteria. Mix by inverting the
tube several times. Leave the tube at
room temperature for 5 minutes.
7) Add 100 µl
(0.1 ml) of potassium acetate (KOAc) to the tube. Mix by shaking the tube
vigorously. You should see a heavy,
clotted precipitate of bacterial genomic DNA, protein, lipid, and carbohydrate.
8) Centrifuge
the tube at full speed for 2 minutes.
9) Remove the
supernatant with a plastic transfer pipet.
Try to avoid the floating white debris and the pellet. Transfer the supernatant to a new,
appropriately-labeled microfuge tube.
10) Add 500
µl (0.5 ml) of isopropanol to the tube containing the supernatant. Mix by inverting the tube several times.
11)
Centrifuge the tube at full speed for 2 minutes.
12) Remove
the supernatant with a plastic transfer pipet, and discard the
supernatant. The pellet contains plasmid
DNA. Resuspend the pellet in 100 µl (0.1
ml) of 50 mM Tris, pH 8.3.
13) Add 50 µl
of 8 M ammonium acetate (NH4Ac).
Mix the contents of the tube, and place it in a dry ice/ethanol bath
until the contents are frozen solid (~10 minutes).
14) Let the
contents of the tube thaw at room temperature, then centrifuge it at full speed
for 2 minutes.
15) The
supernatant contains the plasmid DNA.
Transfer this supernatant to a new, appropriately-labeled microfuge
tube.
16) Add 500
µl of ethanol. Mix the contents by inverting the tube several times. Centrifuge the tube at full speed for 2
minutes.
17) Pour the
ethanol supernatant out of the tube.
Discard the ethanol. Give the
tube a quick spin, then remove the rest of the ethanol from the tube, leaving
the plasmid DNA pellet undisturbed.
18) Allow the
pellet to air-dry for about 5 minutes, then resuspend the pellet in 25 µl of TE
buffer.
Part 2.
PCR Reactions
Procedure
1) Label a microfuge tube with your initials and the letter Q (for “quantification sample”). Dispense 998µl of distilled water into the tube.
2) Using the 2-20µl micropipetter, add 2.0µl of your recombinant plasmid DNA (pick the one that worked best) to the Q tube. Mix well by vortexing.
3) Determine the concentration of your DNA samples using the UV spectrophotometer. Your instructor will demonstrate.
4) Dilute an aliquot of your plasmid DNA sample to a concentration of 4ng/µl.
5) Add 20 µl (= 80 ng) of recombinant plasmid DNA to the tube containing PCR reaction mix (Reaction buffer, Deoxynucleotide triphosphates [dNTPs], Primers, and Taq DNA polymerase), and place it in the thermocycler. When everyone’s samples are in the thermocycler, it will be turned on and run for the proper number of cycles. We will use two different sets of primers, so that each lab pair will perform one PCR reaction with primer pair “A” and one with primer pair “B”.
After the PCR reactions are completed, your instructor will shut down the thermocycler and store your samples for you until next week.
___
Lab
Session 6
1. Run a gel
on the PCR products and isolate the probe DNA fragment.
In this lab
session, you will analyze your PCR products by electrophoresing them on a
special low-melting temperature agarose gel.
From the results of this experiment, you will be able to determine the
size of amplified P-element DNA fragment that resulted from your PCR
reaction. You will isolate the fragments
that contain pure P-element DNA by cutting them out of the gel with a razor
blade. In doing this, you will prepare
pure P-element probe DNA.
Part 1.
Electrophoretic analysis of PCR products