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

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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, Griffiths, A.J.F., W.M. Gelbart,  J.H. Miller, and R.C. Lewontin (1999). Modern Genetic Analysis. New York. W.H. Freeman and Co.  Some non-eukaryotic vectors that are used commonly include:

 

· 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.

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.

 

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

 

Procedure

1) Remove the tubes containing your PCR reaction products from the freezer, and heat them in a 70oC water bath for 5 minutes, then place them on ice.

 

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 tubes containing your PCR products.  Tap the tubes 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) Load load the entire contents of the "l-Hind III" tube,  and 25 µl of each PCR reaction into the sample wells of an agarose gel as shown:

 

Sample Well                                                        

                                                                Number (Starting

                                                                at left)                                                       Sample

                                                                1                                                              "l-Hind III"

                                                                2                                                              “A” PCR Reaction Products

One Pair of Students                           3                                                              “B” PCR Reaction Products

_____________________________4_____________________________Leave Empty_____________

                                                                5                                                              "l-Hind III"

                                                                6                                                              “A” PCR Reaction Products

Second Pair of Students                     7                                                              “B” PCR Reaction Products                                                                                              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 100 volts. Electrophorese until the bromphenol blue (dye) in the samples has migrated to within 5 mm of the positive (red) electrode end of the gel. At 100 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.

 

9) Using information in the “Interpretation of Electrophoresis Data” part of this manual (pp. 20-21), and the guidance of your lab instructor, detemine the size(s)  of the DNA fragments in any bands in the “PCR Products” lanes of your gel.

 

10) Consult the sequence of the P-element and decide which bands on the gel contain fragments of pure P-element DNA.  Your instructor will demonstrate how to cut these bands out of the gel using a clean razor blade, thus isolating pure P-element DNA for use as a hybridization probe for the Southern blot.  After cutting out the appropriate probe DNA bands, place them in a microfuge tube and store the tube in the refrigerator.

 

 


Lab Session 7

1.  Purification of PCR product

2.  Set up the labeling reaction

3.  Prepare the blot for hybridization

4.  Start the hybridization

 

In order to successfully label your probe, you will need to purify your PCR product away from contaminating agarose.  We will do this using a spin columns from a commercially available kit.

Next, you will attach the DIG label to your PCR probe using a process known as random priming. Your instructor will explain how this works

While your labeling reaction is incubating, you can prepare the surface of your Southern blot for the hybridization with the labeled probe,  To do this, you will “pre-hybridize” your blot in the hybridization buffer solution without probe.  This helps reduce background contamination by non-specific binding of probe DNA.

Finally, you will start your hybridization and allow it to incubate overnight.

 

Part 1.  Purification of PCR product

 

Procedure:

1) Obtain a mass for your PCR product: tare the balance with an empty microcentrifuge tube and then weigh your gel slice in its tube.

 

2) Using a micropipetter, add buffer QG into the tube containing the gel slice (add 300ul QG for every 10mg of gel slice), heat gel slice in buffer in a 65oC heat block for 10 minutes or until fully melted.

 

3) Remove gel slice solution from heat block and add  isopropanol (100ul for every 10mg of gel slice).

 

4) Using your micropipetter, transfer gel slice solution to a spin column. Place the spin column in a collection tube and centrifuge at full speed for 1minute (if you have too much volume to fit in the column, fill the column about 2/3, spin this down, discard flow-through.  Then add the rest of your solution to the column and centrifuge a second time).

 

5) Discard the flow through from the collection tube.

 

6) Wash the spin column by adding 500ul of QG buffer to the spin column, and centrifuge at full speed for 1minute.  Discard the flow through from the collection tube.

 

7) Wash the spin column a second time, this time by adding 750ul buffer PE into the spin column, and centrifuge at full speed for 1minute.  Discard the flow through from the collection tube.

 

8) Dry the spin column by spinning an additional minute in the microcentrifuge at 13,000 rpm.

 

9) To elute your DNA, move your spin column to a clean microcentrifuge tube.  Add 50ul of buffer TE to your column, allow the buffer to penetrate the column matrix for at least 1 minute.

 

10)  Elute by centrifuging at full speed for 1minute.  Your DNA will now be in the bottom of the microcentrifuge tube.

 

11) Determine the concentration of your DNA samples using the UV spectrophotometer as you did in session 5.

 

 

Part 2. Set up the labeling reaction.

.

Procedure

1) Label a microfuge tube with your initials and the letter L (for “labeled sample”).  Add 1ug of your PCR probe and bring the volume up to 16ul with sterile distilled water.

 

2) Denature your probe DNA by placing it in a boiling water bath for 10 minutes and then quickly chilling on ice.

 

3) Mix “DIG-High prime” solution with the vortexer, centrifuge briefly.  Using a 2-20ul micropipetter, add 4ul of this solution to your PCR probe. Centrifuge this mixture briefly and incubate at 37oC for one hour.

 

Go to part 3 to prepare your blot while your labeling reaction is incubating.

 

4) After one hour, stop your labeling reaction by adding 2ul of 0.2M EDTA (pH 6.0).  Place your labeling reaction on ice.

 

 

Part 3.  Prepare the blot for hybridization.

 

Procedure:

1) Determine the area in cm of your blot.  Prepare 10mL of “Easy-hyb” hybridization buffer solution for every 100cm2 of blot area. 

 

2) Place your blot in the hybridization tube.  *Pay attention to which side of your blotis which: you want the DNA side facing the interior of the tube.*  Label your tube with your name using tape.

 

3) Add the appropriate volume of hybridization buffer solution.  Use a pipet or forceps to get rid of any bubbles between your blots and the sides of the tube.

 

3) Place your labeled tube in the hybridization oven that has been  pre-heated to 42 oC.  These tubes should be balanced so be sure to put your tube in the oven at the same time as another group so you can balance each other’s tubes.

 

4) Pre-hybridize your blot for at least 30 minutes,  Leave it in the oven until you are ready to hybridize.

 

5) Prepare an additional 3.5mL of hybridization solution buffer for every 100cm2 of blot area in a conical tube.  Pre-heat this in a 42oC water bath until you are ready to start your hybridization.

 

Return to part 2: to stop your labeling reaction.

 

Part 4.  Start the hybridization

 

Procedure:

1) After your probe has pre-hybridized for at least 30 minutes, denature your labeled probe by placing it in a boiling water bath for 5 minutes and then quickly chilling on ice.

 

2) Add the chilled probe to the preheated hybridization buffer solution that you set aside in step5 above.

 

3) Recover your hybridization tube from the hybridization oven.  Discard the prehybridization solution and replace it with the solution containing your labeled probe.

 

4) Place your labeled tube in the hybridization oven, balance as before.  The hybridization will run overnight.

 

 


Lab Session 8

1. Antibody hybridization of  blot

2. Develop blot with chemiluminescent reagent

3. Expose blot to film

 

Last week your lab instructor removed your blot from hybridization and washed it twice with each of two buffer solutions designed to wash away any probe that had stuck non-specifically to the blot while not affecting any probe that was specifically bound to your genomic DNA.  Your blot was then sealed in “washing buffer” solution to store until this week.  Now, your PCR probe should be bound to any P-element sequences in your genomic DNA on the blot.  In order to detect the probe, we will first expose the blot to an Anti-DIG antibody that will bind DIG-labeled probe with high affinity.  This antibody is also conjugated to an alkaline phosphatase (AP) moiety, which we will use in Part 2 to detect the antibody.

The AP moiety on the antibody will, when presented with the appropriate substrate, generate a luminescent product that can be detected on autoradiography film.  The substrate we use is called CSPD.  After incubation with the substrate, you will expose your blot to film and develop the film to see the results.

Today’s lab has many steps, and it is important to work efficiently if you want to be done in three hours.

 

Part 1. Antibody hybridization of  blot

 

Procedure

1) First you will “block” or pre-hybridize your blot,as you did before the labeling hybridization.  This will reduce background contamination from non-specific binding of the antibody. 

 

2)  Prepare a 1X working concentration of blocking solution by diluting 10X blocking solution 1:10 with maleic acid buffer solution.  You will need 50mL blocking solution for every 100cm2 of blot area.

 

2)  Place your blot in the hybridization tube, DNA side facing the interior of the tube.  Label your tube with your name using tape.

 

3) Add the appropriate volume of 1X blocking solution and incubate at 370C for 20 minutes.

 

4) Prepare the antibody solution: you will need 20mL 1X blocking solution for every 100cm2 of blot area. Pre-warm this solution to 37oC

 

5) Your lab instructor will centrifuge the Anti-Digoxigenin-AP solution for 5 minutes at 10,000 rpm and give you an aliquot of the antibody to add the the solution you prepared in step 4.

 

6) Recover your hybridization tube and discard the used blocking solution.   Add your antibody solution and hybridize

By incubating at 37oC for 20 minutes.

 

7) Wash away unbound antibody using the washing buffer solution.  You will perform three short washes of 5 minutes each.  For each wash, discard the solution in the hybridization tube, replace it with washing buffer solution (50 mL for every 100cm2 of blot area).  Place the tube in the hybridization oven for five minutes.

 

Part 2. Develop blot with chemiluminescent reagent

 

Procedure:

1) Remove your blot from the hybridization tube and place in a dish.  Discard the washing buffer solution and equilibrate with detection buffer (20mL for every 100cm2 of blot area).  Allow the pH to equilibrate 2-5 minutes.

 

2) Place your blot with the DNA side facing up on a piece of plastic wrap.  Apply 1mL of CSPD reagent.  *Immediately* cover the blot with a stiff plastic sheet to spread the substrate evenly, and without air bubbles, over the blot.  Incubate for 5 minutes at room temperature.

 

3) Squeeze out excess liquid (but not completely!- it is important for the blot to stay damp though the exposure step) and seal the edges of the plastic wrap around the blot.

 

4) Incubate the damp membrane for 10 minutes at 37oC to enhance the luminescent reaction.

 

Part3.  Expose the blot to film

 

Procedure:

1)  Expose the damp blot to autoradiographic film for 30 minutes. 

2)  Develop film in dark room

3)  If the signal is faint then re-expose the blot to a new piece of film from one hour up to overnight.

 

 

Lab report discussion:  Interpretation of Southern Blot Hybridization Analysis Data

 

1) Obtain the autoradiogram of your hybridized Southern blot. Using information in the “Interpretation of Electrophoresis Data” part of this manual (pp. 20-21), and the guidance of your lab instructor, detemine the sizes of the DNA fragments to which the P-element probe hybridized on your Southern blot.  Did you detect P-elements in the genome of this strain of D. melanogaster?  If so, how many?

 

 

Interpretation of Electrophoresis Data

 

 

1) Figure 7 shows the lengths (in kb) of the DNA fragments in each band resulting from the electrophoresis of l DNA cut with Hind III alone (the DNA size-standards in this experiment).  Examine the bands in the l- Hind III lane of your gel, and determine the lengths of the fragments that make-up each band.  For example, the band that is closest to the sample well represents the largest fragment, which is 23.1 kb long.  Determine the distance (in mm) that each of these l- Hind III bands has migrated from the sample wells.  Using semi-log graph paper,  plot migration distance (in mm) of each l- Hind III band on the X-axis, and DNA fragment length (in kb) of that band on the Y-axis.  Draw a line though the points on your graph.  From this graph, determine the length of the fragments that make up each band in the experimental lanes of your gel  or autoradiogram based on their migration distances.  Do this as follows for each band:

 

                a) Find the migration distance of the band on the X-axis of your graph.

                b) Find the point on the line that is directly above this X-coordinate.

c) Find the point on the Y-axis that is directly to the left of this point on the line.  This Y-coordinate is the size of the fragment (in kb).

 



Project 2:  Isolation and Characterization of Mutations in Drosophila melanogaster

 

 

Introduction

 

One of the most widely used organisms in genetic studies is the fruit fly, Drosophila melanogaster.  Thomas Hunt Morgan pioneered genetic studies with Drosophila at Columbia University in 1911.  Today, there is a large body of knowledge regarding the genetics of the fruit fly.  Drosophila has four pairs of chromosomes (a relatively small number) that have been characterized extensively.  In less than two weeks, the fruit fly undergoes a specific developmental sequence from fertilized egg to larva, prepupa, pupa, then adult.  The geneticist can follow the inheritance of morphological, physiological, and developmental patterns.  In this project, you (the geneticist) will be performing a screen to identify and characterize Drosophila mutants that exhibit morphological abnormalities.  You will then characterize the mutations that you identify, in the hope of learning something about the genes involved.

 

The chromosomes of Drosophila melanogaster

The individual Drosophila has four pairs of chromosomes.  A female has two each of chromosomes 1 (more commonly called the X chromosome), 2, 3, and 4.  A male has one X chromosome, one Y chromosome, and two each of chromosomes 2, 3, and 4.  The Y chromosome and chromosome 4 are both very small, and carry few genes.  The majority of the fly's genes are carried on chromosomes X, 2, and 3.  The X and Y chromosomes are involved in sex determination, and are thus called the sex chromosomes.  Chromosomes 2, 3, and 4 are called the autosomes.  In the fruit fly, sex is determined by the relative number of X chromosomes and autosomes.  If a fly has two X chromosomes, and two of each autosome (an X:autosome ratio of 1:1), it will develop as a female.  If a fly has only one X chromosome, and two of each autosome (an X:autosome ratio of 1:2), it will develop as a male.  In Drosophila, the Y chromosome does not determine maleness! (This is in contrast to the case in mammals, in which the presence of the Y chromosome determines maleness.).  In fact, a fly that has two X chromosomes and a Y chromosome will develop as a female.

 

Development of Drosophila melanogaster

Mating in the fruit fly occurs 6-8 hours after the adult female emerges from her pupal case.  Eggs may be laid at this time, or retained and laid later.  A female receives about 4000 sperm from a male, and stores them in special sacs.  The sperm are released gradually as the eggs are produced.  Each female can lay several hundred fertilized eggs on the surface of a food source.  Each fertilized egg develops over a period of 24 hours into a larva.  The larva burrows into the food source, and eats yeast cells.  Four to five days and two molts (shedding of the larva's exterior cuticle) later, the larva climbs onto a solid surface and pupariates to form a prepupa, which covers itself in a hard pupal case. The prepupa develops into a pupa in 12 hours.  Over the next 4-5 days, the pupa develops into an adult, which emerges from the pupal case in the process of eclosion.  Initially the fly is long and thin, with folded-up wings, and is light in color.  Gradually, the wings expand and the fly takes on a more rounded form and darker color.  The entire life cycle, which takes 10-14 days at 25˚ C, is illustrated in figure 1.

 


 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

                                      Figure 1. The Drosophila life cycle


The study of mutations in Drosophila melanogaster

Mutations are powerful tools in genetic analysis.  The logic behind mutational analysis is that we can learn about the function of a gene by examining what goes wrong when that gene doesn't function properly.  Genes can be altered from their wild-type forms by mutations, which often disrupt or completely eliminate gene function.  Mutational analysis has been called "genetic dissection".  The first stage in a genetic dissection is the hunt for mutants (individual organisms that carry mutant genes).  Mutants occur spontaneously in any population at low frequency.  Through the use of mutagens, however, we can dramatically increase the likelihood of finding useful mutants.  The use of a mutagen to induce mutations is called mutagenesis.  A mutagen that has proven extremely effective in inducing mutations in Drosophila is the chemical, ethyl methane sulfonate (EMS).  EMS can add an ethyl group (-CH2CH3) to many positions on all four bases found in DNA, altering their pairing properties.  The most common change induced by EMS is the addition of an ethyl group to guanine (G), enabling it to pair with thymine (T).  This illegitimate pairing leads to GC --> AT transitions at the next round of replication (see Griffiths textbook, p. 596).

            EMS induces a high proportion of point mutations.  This mutagen is easily administered to adult flies by placing them on filter paper saturated with a mixed aqueous solution of EMS and sugar.  Males fed 0.025 M EMS produce sperm carrying lethal mutations on 70% of all X chromosomes, and on almost every chromosome 2 and 3.  At these levels of mutation induction, it is feasible to look for (screen) for mutations at specific loci, or for mutants that display unusual phenotypes.  In this project, you will be screening flies that have been mutagenized with EMS, to find mutants with abnormal phenotypes.

 

The use of the attached-X chromosome in Drosophila mutagenesis

The majority of interesting mutations that will be detected in a screen following EMS mutagenesis will be recessive, thus yielding no mutant phenotype unless homozygous.  An exception to this requirement of homozygosity for the expression of recessive mutant phenotypes is the case of X chromosome-linked mutations in male flies.  Since a male fly has only one X chromosome (and is thus called hemizygous for all X-linked genes), he will express the abnormal phenotype associated with any recessive mutation on the X.  By making use of a special chromosome called an attached-X chromosome (represented as X^X), it is possible to screen F1 generation males for interesting mutations on the X chromosome.  The attached-X chromosome is a compound chromosome formed by the fusion of two X chromosomes.  An X^X chromosome is inherited as a single unit, and flies carrying an X^X chromosome will be female.  A female that has an X^X chromosome, and also has a Y chromosome can be mated with a normal male to produce X^X/Y female, and X/Y male F1 offspring, as shown in figure 2.  Attached-X chromosomes make it possible to perform a rapid EMS mutagenesis and mutant screen by feeding EMS to Parental generation males, mating them to X^X females, and screening the resulting F1 progeny males for abnormal phenotypes.  A scheme for performing this type of mutagenesis and screen is outlined in figure 3.  In the F1 generation, only the males will carry mutagenized X chromosomes.  Since males are hemizygous for all X-linked genes, recessive X-linked mutations will be expressed in these F1 males.


 

 

 

 

 

 

 

 

 

 

 

 

 

 


 


 

 

 

 

 

 

 

 



This laboratory project

 

In this laboratory project, you will be receiving a large group of F1 offspring resulting from the mating of EMS-treated wild-type males with attached-X females.  This group of F1 flies should contain a number of X-linked mutants.  You are on a search for interesting mutants.  You will be working in pairs, but the entire class will cooperate in this search for mutants, and you can share mutants with each other.  You will discover these mutants by looking for flies displaying phenotypes that vary from wild-type.  An example of such a phenotype would be eyes that are a different color from the wild-type, dark red eye color.  You may also find flies that have abnormal-looking wings, bristles, or other body parts.  Once you have identified as many interesting mutants as you can, you will want to characterize them.  For a given mutation, you will want to determine if that mutation can be transmitted to the next generation.  You also want to determine if each mutation is, in fact, on the X chromosome.  You should design experiments to answer these questions.  Once you have determined which of your mutations are transmissible, it is time to choose one to examine in more detail.  You will want to map the mutation to a specific region of the X chromosome.  This laboratory manual is designed to provide you with guidance.  It is not meant to lead you step-by-step through each experiment.  You should perform this analysis as independently as possible!  With the help of your instructors, you should design and carry out experiments to address the questions listed above.  A potential schedule for this project is outlined below.

 

 

Experimental Procedures (work in groups of 4)

 

 

Lab Session 1

1. Examination of wild-type fruit flies

2. Screening for mutants

 

During this lab session, you will begin screening for mutant flies.  You will first anesthetize and examine a group of wild-type Drosophila.  You should familiarize yourself with the appearance of a wild-type fly, and learn to distinguish males from females.  Once you feel comfortable working with flies, you will be given a culture bottle containing F1 progeny of EMS-treated wild-type males mated with attached-X females.  The bottle will contain attached-X F1 females, and X*/Y F1 males. (See Figure 3)  You will be concentrating on the males, looking for flies with abnormal phenotypes.

 

You will be provided with:

 

a vial containing wild-type fruit flies

a fly anesthetizer

fly anesthetic chemical

a white paper card

a paint brush

a dissecting microscope

a large group of F1 offspring resulting from the mating of EMS-treated wild-type males with

attached-X females.

a vial of virgin attached-X females [C(1)A, y]

empty fly culture vials


 

 

Part 1. Examination of wild-type fruit flies

 

You will first be provided with a vial containing wild-type fruit flies.  Anesthetize all of the flies in the vial as described below, and as demonstrated by your lab instructor.

 

1) Remove the bottom cap of your fly anesthetizer, and take out the foam rubber pad found inside the apparatus.

 

2) "Charge" your fly anesthetizer by putting about 1 ml of Fly Nap anesthetic on the foam rubber pad, and placing the pad back inside the apparatus.  Put the bottom cap back on.

 

3) Remove the top cap from your fly anesthetizer.  Tap the bottom of the vial of flies lightly and rapidly on a pad on your bench, then remove the plug from the vial.  Quickly invert the fly vial over the top of the open anesthetizer, and tap the whole thing lightly and rapidly on a pad, so that the flies fall into the anesthetizer.

 

4) Quickly cap the anesthetizer, and keep the flies in the anesthesia chamber until they all stop moving (this should take a couple of minutes).

 

5) Dump the anesthetized flies out of the anesthetizer onto a white paper card, and view them using a dissecting microscope.

 

6) Practice moving the flies around on the white paper card with a fine paint brush.  Notice the wild-type, dark red color of the eyes.

 

7) Using the diagrams in Figure 4 and provided in the lab room, separate the males from the females.  Males have narrower abdomens than females, and the posterior end of the male abdomen is more darkly pigmented than that of the female.  Males have dark genitalia on the extreme posterior ends of their abdomens that females lack.  Males also have specialized bristles called "sex combs" on their most anterior pair of legs.  If you are having trouble telling males from females by looking at the end of the abdomen, the sex combs will positively-identify a male.

 

8) Once you feel comfortable working with flies and telling males apart from females, it is time to begin screening for mutant flies!

 

 

 

 

 

 

Figure 4. Distinguishing male from female Drosophila

Part 2. Screening for mutants

 

You will be given a large group of F1 offspring resulting from the mating of EMS-treated wild-type males with attached-X females. 

 

1) Anesthetize and examine the flies carefully, concentrating on the males.  The males and females should be easy to tell apart, because the attached-X females have yellow bodies, while the males, unless altered by a newly-induced mutation affecting body color, will have dark tan bodies.  The females will have yellow bodies, because the attached-X chromosomes carry the mutation, yellow (abbreviated, y).  The name of the attached-X chromosome that these females carry is C(1)A, y.  This stands for "Compound Chromosome 1 (the X) of Armentrout (the scientist who made the chromosome), carrying the mutation, yellow".  Both of the X chromosomes that make up C(1)A, y carry the yellow mutation, so the females are homozygous for this recessive mutation, and display the mutant yellow body color.  Keep in mind that, if you see a yellow-colored male, it is potentially due to a newly induced X-linked mutation, and you should examine it further. 

 

2) Look for males that show any differences from wild-type.  Save each mutant-looking male that you find in his own culture vial.  If you absolutely cannot find a mutant, DON'T WORRY!  This will be a team effort, and you can get a mutant from another group of students if you need to.  Your instructor will also provide additional mutants.

 

3) With the help of your lab instructor (if you want it) design and begin a experiments to determine if the mutations you have identified are transmissible to the next generation.  You’ll be provided with everything (including flies) that you need to set up these experiments.  You should discuss your approach with your instructor.  You should also be thinking about how you will map your mutations to a region of the X chromosome.

 

 

Lab Session 2

 

During this lab period, you will interpret the results of  your experiment designed to determine if each of your newly identified mutations is transmissible. 

 

1) Anesthetize the flies in the vials that you set up crosses in during Lab Session 1.  Examine all of the flies.  These flies, the progeny of the cross(es) you set up during Lab Session 1, are the F2 generation.  Look for F2 flies displaying the mutant phenotypes you identified during Lab Session 1. Pay attention to differences between males and females.

 

-Which of your mutations are transmissible?

 

-Of the mutations that are transmissible, can you tell if any are dominant or recessive? Can you tell if any are X-linked or autosomal? Explain your reasoning.

 

-If a particular mutation did not transmit, think about why that may be. 

 

2) If a particular mutation is transmissible, begin your experiment to map it to a region of the X- chromosome.  You will be provided with virgin female fruit flies homozygous for a multiply-marked X-chromosome.  This multiply-marked X-chromosome carries several different recessive mutations that are easy to identify.  It is NOT an attached-X chromosome!  An example of a multiply-marked X-chromosome is the  y   cv  v  f   chromosome.  This chromosome carries recessive mutant alleles of four genes that are spaced along the length of the chromosome.  These genes are:

 

y = yellow    (maps to the telomere, or extreme left end of the X-chromosome, Map Position = 0) Female flies homozygous (or male flies hemizygous) for mutations in yellow have very light yellow body color, as opposed to the tan body color of wild-type flies.

 

cv = crossveinless   (Map Position = 13.7, which means it is 13.7 Map Units to the right of the telomere) Female flies homozygous (or male flies hemizygous) for mutations in crossveinless lack a certain set of veins that are supposed to be in their wings.

 

v = vermillion   (Map Position = 33.0, which means it is 33 Map Units to the right of the telomere) Female flies homozygous (or male flies hemizygous) for mutations in vermillion have an abnormal pinkish eye color, as opposed to the dark red eye color of wild-type flies.

 

f = forked   (Map Position = 56.7, which means it is 56.7 Map Units to the right of the telomere) Female flies homozygous (or male flies hemizygous) for mutations in forked have an abnormal sharp bend in the end of their bristles, which makes forked bristles appear quite different from the straight, pointed bristles of wild-type fruit flies.

 

A schematic map of the y  cv  v  f  chromosome would look like this:

 

 

Telomere                                                                  Centromere

 

                                    Map Position

 

0                      13.7                 33.0                 56.7

 

y                      cv                     v                      f________

 

 

 

If the mutation that you wish to map is X-linked, set up a cross of males hemizygous for your newly-identified mutation with virgin females homozygous for the multiply-marked X-chromosome.  You will interpret the results of this experiment during Lab Session 3.

 

 


Lab Session 3

 

During this lab period, you will continue your mapping experiments.  You will interpret the results of the cross(es) that you set up during Lab Session 2, and set  up another cross (or crosses).

 

1) Anesthetize the flies in the vials that you set up crosses in during Lab Session 2.  These flies, the progeny of the crosses you set up during Lab Session 2, are the F3 generation.  Examine all of the flies.  Look for flies displaying the mutant phenotypes you identified during Lab Session 1.  Pay attention to differences between males and females.

 

-What do the males flies look like?

 

-What do the female flies look like?

 

-Do any of the flies display the mutant phenotype that you identified in Lab Session 1?

 

-From your results, can you determine whether your newly-identified mutation is dominant or recessive?

 

-From your results, can you determine whether your newly-identified mutation is allelic to any known mutation?

 

2) The F3 females are heterozygous for  the X-chromosome carrying your newly-identified mutation,  and the multiply-marked X-chromosome.  During meiosis in these females, crossing-over can occur  between these two X-chromosomes, resulting in eggs carrying recombinant X-chromosomes.  By examining the F4 progeny resulting from a cross between these females and appropriate male flies, you can determine the recombination frequencies between your newly-identified mutation and the known mutations on the multiply-marked X-chromosomes.   This will enable you to calculate a map position for your new mutation.

Since you are dealing with X-linked mutations, you can plan to restrict your analysis of the F4 progeny to the F4 males.  Each F4 male will inherit a single X-chromosome from his heterozygous F3 mother, and will display the phenotypes associated with any mutations on that X-chromosome.  Each F4 male will either be parental-type or recombinant with respect to the several mutations you are working with.  Determining the percentage of F4 male progeny that are recombinant will enable you to calculate a map position for your newly-identified mutation. 

Since you will be looking at only the males in the F4 generation, the genotype of the males that you cross with your F3 heterozygous females does not matter. You can consider these males to be merely sperm donors.  Set up a cross(es) of your F3 heterozygous females with available males.        

 

 


Lab Session 4

 

During this lab period, you should be able to finish your proposed mapping experiment, and determine what region of the X chromosome each of your mutations maps to. 

 

1) Anesthetize the flies in the vials that you set up crosses in during Lab Session 3.  These flies, the progeny of the crosses you set up during Lab Session 3, are the F4 generation.  Examine all of the flies.  Separate the males from the females, and discard the females.  You will focus this analysis on the F4 males only. 

 

2) Pick three mutations to focus your attention on .  These three mutations must include your newly-isolated mutation and two of the mutations on the multiply-marked X-chromosome.  You can now consider your study a three-point cross, as described on pages 156-167 of the textbook Griffiths et al. (2002) Modern Genetic Analysis, 2nd Edition. New York, W.H. Freeman and Company.  Your first job will be to determine the phenotype (mutant or wild-type) of each of the F4 males with respect to the three mutations you are considering.  You can then figure out the numbers of F4 males that are parental-type and recombinant-type with respect to each of the three mutations.

 

3) After scoring all of the F4 males for phenotype, determine which class of progeny represent the double cross-overs.  Determine the number of single cross-overs that occurred between each of the three mutations.  With these numbers, you should be able to map the three mutations relative to one another.  Since you know the map positions of two of the mutations, you should be able to determine a map position for your newly-isolated mutation. 

 

4) Draw a map of the X-chromosome showing the map positions of your newly-isolated mutation and the two other mutations you used.