Lectures 4 + 5: Lecture Notes + Textbook
Clone: - Identical copy of something (ie exact copy of DNA)
- Needed for sequencing
- Bacteria reproduce by cloning themselves
- One example of synthetic cloning is PCR
The basic tools of gene exploration include
a) Restriction enzyme analysis
b) Blotting techniques (ie Southern blotting for DNA, Northern blotting for RNA)
c) DNA sequencing
d) PCR (polymerase chain reaction)
Restriction Enzymes
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Also known as restriction endonucleases
Discovered by Arber and Smith
Nathans pioneered their use in the late 1960s
Recognize specific base sequences in double helical DNA, and cleave, at specific sites, both strands of a duplex
containing the recognized sequences
Extremely important – used in analyzing chromosome structure, sequencing DNA, isolating genes, and
creating new DNA for cloning
Restriction enzymes are naturally found in a wide range of prokaryotes
They serve to cleave foreign DNA – the organism’s own DNA is not cleaved by its own restriction enzymes
though because the sites recognized by the restriction enzymes are protected through methylation (only on
own DNA)
Many restriction enzymes recognize specific sequences of four to eight base pairs
They act through hydrolyzing the phosphodiester bond in each strand
Usually, rotational /bilateral symmetry – that is, the recognized site is palindromic (inverted repeat)
For example, in the top picture to the right:
3’ G A A T T C 5’
Notice how the top strand and bottom one are inverses of each other
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The ◊ represents axis of symmetry (horizontal)
5’ C T T A A G 3’
The restriction enzyme will usually cut somewhere in the shaded area
It always cuts the bond on the 3’ side of the axis of symmetry
Thus, if this enzyme hydrolyzes the bond between the G and A, the
3’ A A T T C 5’
newly cut enzyme will look like:
◊
(It only hydrolyzes the 3’ region of each single strand)
5’ C T T A A 3’
Notice the staggered DNA that is created – these are sticky ends
(more on that later)
Restriction Enzymes usually recognize short base sequences
(short segments = higher probability of that sequence appearing multiple times = more clones
long segments = lower probability of that sequence appearing multiple times = less clones)
Southern Blotting
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Used to identify a specific sequence located within a DNA fragment
First, a mixture of restriction fragments is separated by electrophoresis through agarose gel
It is then denatured to form single stranded DNA, and transferred to a nitrocellulose sheet
The positions of the DNA fragments are preserved on the nitrocellulose sheet
These single stranded DNA are then exposed to a 32P labeled probe (radioactive probe)
The probe hybridizes with restriction fragments having its complementary sequence
Autoradiography displays position of this probe-fragment duplex
RNA blot is called a Northern Blot, protein blots are called Western Blots
DNA Sequencing – the Sanger Dideoxy Method
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Developed by Sanger and co
Works through controlled interruption of enzyme replication
This procedure is performed by four reaction mixtures at the same time
In all of these mixtures, DNA polymerase is used to make the complement of a particular sequence
It is primed by a fragment that contains the complementary sequence to a known sequence
What makes this method special is its use of specific “terminators” – each reaction contains a small amount
of a 2’-3’ –dideoxy analog of one of the nucleotides (a different nucleotide for each reaction mixture –
remember, there are 4 reaction mixtures, so 1 mixture will have the adenine form of the dideoxy analog,
another will have the guanine form, and the other two will have thymine and cytosine)
Normally, the polymerase adds a complementary base, then moves to the next nucleotide and does the same
However, in addition to the regular nucleotides that the DNA polymerase usually add, they can also add these
2’-3’ –dideoxy analogs, as long as the base is the same. For example, if it needs to add an adenine to
complement a thymine, it can also add a 2’-3’ – dideoxy analog of adenine (but it can not add a dideoxy
version of guanine to complement thymine)
However, if the dideoxy analog version of the nucleotide base is added instead of the pure nucleotide base,
the polymerase is no longer able to add a nucleotide after it (this phenomenon is known as chain
termination). The reason is that this dideoxy analog version of the nucleotide base does not have the OH
group at the 3’ end, and thus, polymerase can no longer add bases
The concentration of this dideoxy analog is low enough though that this “chain termination” will only take
place occasionally (ie most of the times polymerase will add the regular nucleotide, but sometimes it will add
this dideoxy version)
Thus, different length fragments will be produced, but all of them will be terminated by a dideoxy analog
base, and as such, these fragments of different length will correspond to the positions of its complement base
on the regular DNA (sorry, I probably made this confusing – for example, the dideoxy analog dGTP (guanine
base) would correspond to where a C would occur in the DNA sequence)
Four sets (one for each base) of these chain terminated fragments then undergo electrophoresis, and the base
sequence of the new DNA is read from the autoradiogram of the four lanes (remember, this is the new DNA
– the sequence of the old DNA would be the complement to this new DNA)
Fluorescent tags can also be used – it can either be attached to the priming fragment (a different color for
each of the four reaction mixtures), or to the dideoxy analog itself (a different color for each of the four
analogs)
If added to the priming fragment, the four reactions are combined, then subjected to electrophoresis, and the
separated bands of the DNA are detected by their fluorescence as they emerge from the gel
If added to the dideoxy analog, only 1 reaction mixture is required
Fluorescent labels are considered good because it eliminates the use of radioactive reagents and can be readily
automated.
Sorry, I know I made that a little more painful – the best advice I can actually give is to go to the biochemistry website
(www.whfreeman.com/biochem5, click on the animated techniques section, and click on chapter 6, dideoxy
sequencing of DNA – after watching that (it’s a quicktime video), everything should be crystal clear)
DNA Amplification – The Polymerase Chain Reaction
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Founded by Kary Mullis in 1984
Method for cloning a sequence within a long strand of DNA
The sequence that is to be copied doesn’t need to be known – only the sequences flanking the target
sequence must be known (so that a proper primer can be made and attached to that site)
PCR is carried out in cycles, with exponential growth after each cycle
In preparation, one prepares a tube that contains the template DNA, a set of primers that flank the target
DNA, Taq Polymerase (Taq polymerase is used because it is heat resistant), nucleotides (lots of them so you
don’t run out), and magnesium (Dr. Hampson said polymerase requires magnesium to do its job)
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Step 1 – Strand separation: The solution is heated to 95ºC
to separate the parent DNA molecule into 2 strands
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Step 2 – Hybridization of Primers: The solution is
cooled to 54ºC to allow the primer to hybridize
(anneal) to the DNA strand. One primer hybridizes
to the 3’ end of one strand, the other primer
hybridizes to the 3’ end of the other strand. Primers
are typically 20-30 nucleotide bases long, and parent
DNA duplexes do not form because there is an
excess of primers (that is, a single strand of DNA
does not bind with its complement because the
primer “beats it to the punch”
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Step 3 – DNA Synthesis: The solution is heated to
72ºC (optimal temperature for Taq polymerase),
and the polymerase elongates both primers in the
direction of the target sequence (remember, DNA
synthesis always takes place in the 5’ to 3’
direction). Only after the third cycle is a double
stranded DNA of the target sequence produced,
and after the third cycle, these short strands, which
represent the target sequence, are exponentially
amplified
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These three steps can be carried out repeatedly by changing the temperature of the mixture
No reagents need to be added after the first cycle
Amplification is one millionfold after 20 cycles, and 1 billionfold after 30 cycles
Remember, the sequence to be amplified does not need to be known – only flanking sequences
The target sequence can be much larger than the primers (Targets > 10kb have been amplified)
Primers do not have to be perfectly matched to flanking sequences to amplify targets. With the use of
primers derived from a gene of known sequence, it is possible to search for variations on the theme.
Families of genes are being discovered this way by PCR (I don’t really get this point, but it’s in the text)
PCR is highly specific because of the stringency of hybridization at high temperatures. Stringency refers to
specificity/exactness, and the required closeness of the match between primer and target can be controlled by
temperature and salt.
PCR is exquisitely sensitive – a single DNA molecule can be amplified and detected
PCR is also a powerful technique in medical diagnostics, forensics, and molecular evolution as it provides a
means of quick amplification (which then can be used to look for specific things, such as polymorphisms, or
parentage (through amplification of different genes)
The Formation of Recombinant DNA Molecules – Restriction Enzymes and DNA Ligase as Tools
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A DNA fragment of interest is covalently joined to a DNA vector
One of the most important properties of a vector is that it can replicate autonomously (Dr. Hampson uses
the term epichromosomally) in an appropriate host – that is, vector DNA replicates independently of host DNA
The vector is prepared for accepting a new DNA fragment by cleaving it at a single specific site with a
restriction enzyme. These restriction enzymes usually produce staggered cuts (complementary single stranded
ends) – these are known as cohesive or sticky ends because they have a specific affinity for each other
Thus, any DNA fragment can be inserted into this plasmid if it has the same sticky ends, and this is simply
done by taking a larger piece of DNA and applying the same restriction enzyme to it as was applied to the
vector (plasmid) DNA
The DNA is then inserted into the plasmid vector and DNA ligase joins the two together, forming a
recombinant DNA molecule. The chains joined by ligase must be in a double helix, and an energy source (ie
ATP, NAD+) is required
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The use of a short, chemically synthesized DNA
linker – first, the linker is covalently joined to the
ends of a DNA fragment or vector. In this
example, the 5’ end of a decameric linker and a
DNA molecule are phosphorylated and joined by
ligase from T4 phage
This ligase forms a covalent bond between bluntended (flush ended / non sticky-end) double-helical
DNA molecules
Cohesive ends are produced when these terminal
extensions are cut by an appropriate restriction
enzyme
Thus, sticky ends corresponding to a particular
restriction enzyme can be added to virtually any
DNA molecule (because flush end is ligated)
Therefore, when using a linker, the linker itself is cleaved by the restriction enzyme (the DNA fragment is
not). Otherwise, the restriction enzyme is applied to the DNA fragment itself
Types of Vectors
Plasmids
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circular, duplex DNA molecules
occur naturally in some bacteria
range in size from 2 to several hundred kilobases (Dr. Hampson said that ideal size is between 5-10 kilobases)
they can carry genes for inactivation of antibiotics, production of toxins, and the breakdown of natural
products
they are accessory chromosomes, and can replicate independently of the host chromosome – in Dr.
Hampson’s terms, they reproduce epichromosomally
Not all bacteria have plasmids, but some can house up to 20 of them
An example of a plasmid is the pBR322 plasmid, which is very useful in cloning
pBR322 contains genes for resistance to tetracycline and ampicillin
different restriction enzymes can cleave pBR322 at a variety of unique sites at which DNA fragments can be
inserted
Insertion of DNA at EcoRI restriction site does not alter any genes for antibiotic resistance
Insertion at the HindIII location, however, inactivates the gene for tetracycline resistance – this is an effect
known as insertional inactivation. Cells containing pBR322 with a DNA insert at this restriction site are resistant
to ampicillin still, but are sensitive to tetracycline – thus, they can be readily selected
Cells that failed to take up a vector are sensitive to both tetracycline and ampicillin (since they don’t carry
vector that has antibiotic resistance)
Cells that contain the vector without a DNA insert, or a DNA insert at the EcoRI location are resistant to
both ampicillin and tetracycline
This notion of selectivity is important, as it allows scientists to easily get rid of bacteria cells they do not want
(ie those without recombinant DNA vector) through the use of antibiotics
Lambda () Phage (Bacteriophage)
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bacteriophages are great at infecting bacteria, and the  phage, like most phages, can enjoy 2 possible
lifestyles:
The lytic pathway, where viral functions are fully expressed – viral DNA and proteins are quickly produced
and packaged into virus particles. This leads to the lysis (destruction) of the host cell and the appearance of
about 100 progeny virus particles (virions)
The lysogenic pathway, where the phage DNA becomes inserted into the host-cell genome and can be
replicated with host-cell DNA for many generations
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Mutant  phages have been designed for cloning
– an especially good one is gt –  as it
contains only two EcoRI cleavage sites ( phages
normally have five)
After cleavage, the middle segment of the 
phage can be removed, and the two remaining
DNA fragments (called arms) have a combined
length equal to 72% of the normal genome
length
However, this amount of DNA is too small to
be packaged into a particle because only DNA
measuring from 75% - 105% can be readily
packaged
Thus, inserting a suitably long DNA insert (such
as 10kb) between the two “arms” of the - useful
 DNA will create a suitable length (93% of
normal length) recombinant DNA molecule
This is extremely useful for cloning genes
M13 Phage
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useful vector for cloning DNA, and is especially useful for sequencing
inserted DNA
the single-stranded DNA in the virus particle, called the (+) strand, is
replicated through an intermediate circular double-stranded replicative form
(RF) containing (+) and (-) strands
only the (+) strand is packaged into new virus particles
about 1000 progeny M13 are produced per generation
M13 does not kill its bacterial host, thereby allowing large quantities of M13
to be grown and easily harvested
M13 is prepared for cloning by cutting its circular RF DNA at a single site
with a restriction enzyme
The cut is made in a polylinker region, a region that contains a series of
closesly spaced recognition sites for restriction enzymes (only one of each
such sites is present in the vector)
Foreign DNA (double stranded) fragment produced by cleavage with the
same restriction enzyme is then ligated to cut vector
Foreign DNA can be inserted in two different orientations because the ends
of both DNA molecules are the same
Thus, half the new (+) strands packaged into virus particles will contain one
of the strands of foreign DNA, and half will contain the other strand
DNA cloned by M13 (through infection of E. coli) can be easily sequenced
as an oligonucleotide that hybridizes adjacent to the polylinker region is
used as a primer for sequencing the insert
This oligomer is called a universal sequencing primer because it can be used to
sequence any insert
M13 is ideal for sequencing but not for long-term propagation of
recombinant DNA because inserts greater than 1 kb are not stably
maintained
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another vector type includes phagemids, which are a combination of plasmid and a phage. It is the most
commonly used vector now, and an example of one is a cosmid. Cosmid = phagemid that is a hybrid of a 
phage and a plasmid, and serves as a vector for large DNA inserts – up to 45 kb
other vectors include BACs (Bacterial Artificial Chromosomes) and YACs (Yeast Artificial Chromosomes),
and these are used for very large pieces of DNA
There are 2 important sites in a vector molecule
The first site is the MCS (multiple cloning site). The multiple cloning site is a relatively small piece of DNA
(usually 3 kilobases) that is a series of unique restriction sites. Therefore, when a restriction enzyme cleaves
the molecule, it only cleaves at this one site, and in a plasmid, the circular DNA becomes linearized
The MCS is digested, giving way for foreign DNA to be inserted
The MCS is the only region in the gene that can be cut – this is important because if the plasmid was to be
cut anywhere else, it could disrupt gene function
Secondly, there is a site that codes for antibiotic resistance. All vectors have genes that code for antibiotic
resistance so that when an antibiotic is added, only cells without vector are killed – selection process
Cloning Genes
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DNA to be cloned is inserted into a vector (ie pBR322 plasmid)
Joined together by DNA ligase, forming a recombinant DNA molecule
Introduced into host cells by transformation (permeabilize bacterial cell to let plasmid through) or viral
infection
Select for cells containing a recombinant DNA molecule
Add antibiotics to kill bacteria without recombinant DNA (remember, plasmid codes for antibody resistance)
Also, remember that vector DNA reproduces epichromosomally (independent of bacterial DNA replication)
Bacteria cells are used because they replicate quite quickly, and vector DNA uses bacterial cell machinery
And voila, we have multiple clones of the gene/sequence inserted into the plasmid
Specific Genes Can Be Cloned From Digests Of Genomic DNA
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one can isolate a specific DNA segment several kb long out of a
genome containing more than 3 million kb
step 1: digestion – a restriction enzyme digests many molecules of
total genomic DNA into large fragments
step 2: gel electrophoresis – random population of overlapping DNA
fragments is separated by gel electrophoresis to isolate a set about 15
kb long
step 3: recombination with vector – synthetic linkers are attached to
the ends of these fragments. Restriction enzymes then cleave these
linkers to create sticky ends, and this bulk is then inserted into a
vector (which has been prepared with the same sticky ends)
step 4: E. Coli infection – E. coli bacteria are then infected with these
recombinant phages. The resulting lysate contains fragments of
human DNA housed in sufficiently large number of virus particles to
ensure the whole genome is represented. These phages constitute a
genomic library (all the DNA of an organism broken up into fragments
and repackaged into viruses). As these phages can reproduce
indefinitely, the library can be used repeatedly over long periods of
time
Picture to the left is Figure 6.18 from the text, and it represents the
creation of a genomic library (the genomic library can be created from
a digest of a whole eukaryotic genome)
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step 5: screening – the genomic library is then screened for the very small portion of phages that hold the
gene of interest. However, for a 99% success rate, over 500 000 clones must be screened, and as such, a
rapid method of screening is needed:
1) dilute suspension of recombinant phages is plated on lawn of bacteria. Where each phage particle has
landed and infected a bacteria, a plaque containing identical phages develops on the plate
2) a replica of this master plate is made by applying a sheet of nitrocellulose – infected bacteria and phage
DNA released from lysed cells adhere to the sheet in a pattern corresponding to the plaque. Also, intact
bacteria on this sheet are lysed with NaOH, which also serves to denature the DNA
3) the denaturing of DNA is extremely important, because it allows for the hybridization of the DNA to a
32P-labeled probe, and this probe is complementary to the sequence of interest
4) the presence of a specific DNA sequence in a single spot on the replica can be detected by using a
radioactive complementary DNA or RNAM molecule as a probe. Autoradiography then reveals teh positions
of spots that have the recombinant DNA
5) the corresponding plaques are picked out of the intact master plate and grown
Long Stretches of DNA Can Be Efficiently Analyzed by Chromosome Walking
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 phage vectors only consist of fragments about 15 kb long, yet many eukaryotic genes are much longer (even
2000 kb long!)
thus, it is easier to use cosmids (they can house fragments 45 kb long), or even better, YACs (yeast artificial
chromosomes) or BACs (Bacteria Artificial Chromosomes)
step 1: digestion – genomic DNA is partly digested by a restriction enzyme that cuts, on the average, at
distant ends
step 2: gel electrophoresis – the fragments are separated by gel electrophoresis
step 3: insertion into YAC – the large fragments are eluted and ligated into YACs. Artificial chromosomes
(be it BAC or YAC) bearing inserts from 100 to 1000 kb are efficiently replicated in yeast cells
step 4: chromosome walking – fragments in a cosmid or YAC/BAC library are produced by random
cleavage of many DNA molecules – therefore, there will be overlap in some fragments, and chromosome
walking takes advantage of these overlaps
For example, probe A’ selects its complementary region A via hybridization, and we will assume that this
fragment containing A also contains region B
A new probe B’ can be prepared by cleaving this fragment between regions A and B and subcloning region B
The library is then screened again with probe B’ this time, and new fragments containing region B will be
found. Some of these will contain a previously unknown region C
We can apply the same steps again to get probe C’, and this process can be repeated over and over again
This process of subcloning and rescreening is called chromosome walking, and long stretches of DNA can be
analyzed in this way provided that each of the new probes is complementary to a unique region.
Complementary DNA Prepared from mRNA Can Be Expressed in Host Cells
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mammalian DNA poses a problem in the sense that if it wants to be expressed in bacteria, it needs to lose its
introns as bacteria can not process introns
however, since mRNA has not introns, if a bacteria picks up a recombinant DNA that is complementary to
mRNA, then the protein can be expressed in the bacteria
an example of this is in the production of insulin – DNA that is complementary to mRNA for proinsulin
(precursor of insulin) is joined to a plasmid, which is then inserted into a bacteria. The bacteria is then able to
make proinsulin, and in fact, much of today’s insulin is produced by bacteria
Formation of cDNA (Complementary DNA)
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the enzyme reverse transcriptase is needed
normally, a retrovirus uses this enzyme to form a DNA-RNA hybrid in replicating its genomic RNA
basically, reverse transcriptase synthesizes a DNA strand complementary to an RNA template if it is provided
with a DNA primer that is base-paired to an RNA template (the 3’OH group must be free too)
The oligo(T) primer is used, because it simply attaches to the poly A tail of most mRNA (although most
mRNA have this tail, some don’t – either a universal primer is used, or a random one)
If these conditions are met, reverse transcriptase basically synthesizes the rest of the DNA molecule in
complementary base-pair fashion (as long as it is in the presence of the four deoxyribonucleoside
triphosphates)
Once it is done, a RNA-DNA heteroduplex is formed
The RNA from this heteroduplex is hydrolyzed (degraded) through raising the pH
The DNA is not degraded though because DNA is resistant to alkaline hydrolyisis
The DNA is then treated with the enzyme terminal transferase, which simply adds nucleotides (for example, lots
of Guanine nucleotides) to the 3’ end of the DNA
This is done so a specific primer, oligo(C) if using the poly G tail from above, can bind to the residues and
prime the synthesis of the second DNA strand – this is how double stranded DNA is created from mRNA
Once this is done, synthetic linkers can be added to this double-helical DNA for cleavage and ligation to a
suitable vector
Complementary DNA for all mRNA that a cell contains can be made this way, and inserted into vectors,
which are then inserted to bacteria. Such a collection is called a cDNA library.
Quick note – since cDNA library contains no introns, it is smaller than genomic DNA library. Also, cDNA
is representative from the tissue it came from, whereas genomic DNA reflects the entire organism
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cDNA molecules can be inserted into vectors that favor their
efficient expression in hosts such as E.coli. The vectors that are
able to produce efficient expression of genes are known as
expression vectors
To maximize transcription, the cDNA is inserted into the vector
in the correct reading frame near a strong bacterial promoter site
These vectors also ensure efficient translation by encoding a
ribosome-binding site on the mRNA near the initiation codon
Clones of cDNA can be screened on the basis of their capacity to
direct the synthesis of a foreign protein in bacteria
A radioactive antibody specific for the protein of interest can be
used to identify colonies of bacteria that have the corresponding
cDNA vector
This is done the exact same way as genomic library DNA
screening – spots of bacteria on a replica plate are lysed to release
proteins, which bind to an applied nitrocellulose filter. A 123Ilabeled antibody specifric for the protein of interest is added, and
autoradiography reveals the location of the desired colonies on
the master plate
DNA Fingerprinting
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Done through restriction mapping – in this process, scientists hone in on variable parts of DNA
Step 1: obtain DNA and extract DNA
Step 2: digest DNA with more restriction enzymes
Step 3: separate DNA fragments by gel electrophoresis
Step 4: visualize bands with specific radiolabeled probe or DNA stain (ethidium bromide)
Step 5: Compare banding pattern between 2 or more samples (ie suspect & evidence sample from crime
scene)
Step 6: Determine whether or not there is a match
This works because there is a 0.1% difference in DNA between individuals
Restriction-fragment-length polymorphism (RFLP) analysis in humans examines the 0.1% that is different
Polymorphism just refers to genetic variation within individuals
If polymorphism occurs at restriction enzyme site, restriction enzyme will only cut at a specific site.
Therefore, this whole fingerprinting technique relies on polymorphisms – in comparing 2 samples (ie suspect
and evidence), if the suspect has a polymorphism at the restriction site, the restriction enzyme will not be able
to cut the fragment. The evidence sample, which does not have the polymorphism at the restriction site, is
cut. When run through gel electrophoresis, the banding patterns will be different, and thus, in this case, there
is no match (this is pretty much the basis of forensics and all that stuff you see on CSI)
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Outside the realm of forensics, restriction fragment length polymorphisms also have uses, such as in
diagnosing diseases
For example, take sickle cell anemia:
A normal gene has the code:
while the sickle cell anemia code is:
C C T G A G G
G G A C T C C
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C C T G T G G
G G A C A C C
The normal gene is usually cleaved at 3 sites with a restriction enzyme, but in cells that suffer from sickle cell
anemia, it is only cleaved at two sites
Thus, RFLP provides a good way to diagnose if someone has sickle cell anemia or not
Gene-Expression Levels Can Be Comprehensively Examined
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Most genes are present in the same quantity in every cell (one in haploid cell, two in diploid cell)
The level at which the gene is expressed is indicated by mRNA quantities, and varies from no expression to
hundreds of mRNA copies per cell
Gene expression varies from cell type to cell type (this helps distinguish cells)
Level of gene expression in tissue can be measured through DNA microarrays or gene chips
DNA microarrays are high-density arrays of oligonucleotides that can be:
constructed either through light-directed chemical synthesis carried out with photolithographic
microfabrication techniques (used in semiconductor industry) or
by placing very small dots of oligonucleotides or cDNAs on a solid support such as a microscope slide.
Fluorescently labeled cDNA is hybridized to the chip to reveal the expression level for each gene, identifiable
by its known location on the chip
The intensity of the fluorescent spot on the chip reveals the extent of transcription of a particular gene
New Genes Inserted into Eukaryotic Cells Can Be Efficiently Expressed
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as bacteria are not able to perform post-translational modifications, such as specific cleavage of polypeptides
and the attachment of carbohydrate units, many eukaryotic genes can only be properly expressed in
eukaryotic host cells
Introduction of recombinant DNA molecules into cells of higher organisms can also be a source of insight
into how their genes are organized and expressed
Recombinant DNA molecules can be introduced into eukaryotic cells in different ways
One way is to have foreign DNA molecules precipitated by calcium phosphate, as these are taken up by
animal cells. A small fraction becomes incorporated into chromosomal DNA. While the efficiency of
incorporation is low, the method is still useful because it is easy to apply
Another method has the DNA microinjected into cells. A fine-tipped (0.1 m diameter) glass micropipet
containing a solution of foreign DNA is inserted into a nucleus. In mice, around 2% of injected mice cells
contain the new gene
Another method uses viruses to bring the new genes into animal cells. Retroviruses are the choice vector
because the double helical DNA from its genome, produced by the action of reverse transcriptase, becomes
randomly incorporated into hose chromosomal DNA. This version of viral DNA is called proviral DNA, and
can be efficiently expressed by host cells and replicated along with normal cellular DNA. As a bonus,
retroviruses do not usually kill their hosts, so the expression can be seen over a period of time
2 other viruses used are vaccinia virus, a large DNA-containing virus that replicates in the cytoplasm of
mammalian cells and shuts dwon host-cell protein synthesis, and baculovrius, which infects insect cells, which
can be conveniently cultured. Insect larvae infected with this virus can serve as efficient protein factories
Vectors based on these large-genome viruses have been engineered to express DNA inserts efficiently
Gene Disruption Provides Clues to Gene Function
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A gene’s function can be probed by inactivating the gene and looking for resulting abnormalities. This
process is known as gene disruption (or gene knockout)
This method relies on the process of homologous recombination. Through this process, regions of strong
sequence similarity exchange segments of DNA. Foreign DNA inserted into a cell can therefore disrupt any
gene that is at least in part homologous by exchanging segments
The disrupted gene is then inserted into mice embryos, and mice lacking the gene are produced
An example of gene disruption in mice is the disruption of the gene myogenin. When both copies for
myogenin are disrupted, the animal dies at birth because it lacks functional skeletal muscle
Heterozygous mice containing one copy of the normal gene and one copy of the disrupted gene appear
normal, indicating that the level of gene expression is not essential for its function
Site Specific Mutagenesis
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New genes with designed properties can be constructed by making three kinds of directed changes to the
DNA sequence – deletions, insertions, and substitutions
Substitutions: Oligonucleotide-Directed Mutagenesis
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one nucleotide in a DNA sequence is simply substituted with another – this is usually done to change one
amino acid to another
for example, if we want to change a serine group to a cysteine, all one needs is a plasmid containing the gene
or cDNA for the protein, and one needs to know the base sequence around the site to be altered. If the
serine is coded by TCT, one needs to change the C to a G to get TGT, which codes for cysteine
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This type of mutation is called a point mutation because only one base is altered
The key to this mutation is to prepare an oligonucleotide primer that is complementary to this region of the
gene except for one point – in this case, it is TGT instead of TCT
The two strands of the plasmid are separated, and the primer is then annealed to the complementary strand
The mismatch of 1 base pair of 15 is tolerable if the annealing is carried out at an appropriate temperature
After annealing to the complementary strand, the primer is elongated by DNA polymerase, and the double
stranded circle is closed by adding DNA ligase
Subsequent replication of this duplex yields two kinds of progeny – half with the original TCT sequence, and
half with the mutant TGT sequence
Expression of the new TGT sequence will produce a protein that has cysteine instead of serine at the one site
Insertions: Cassette Mutagenesis
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a plasmid DNA is cut with a pair of restriction enzymes to remove a short segment
a synthetic double-stranded oligonucleotide (the cassette) with cohesive ends that are complementary to the
ends of the cut plasmid is then added and ligated
Each new plasmid now contains the desired mutation
It is convenient to introduce into the plasmid unique restriction sites spaced about 40 nucleotides apart so
that the mutations can be readily made anywhere in the sequence
Deletions
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A specific deletion can be produced by cleaving a plasmid at two sites with a restriction enzyme, then ligating
to form a smaller circle. This approach usually removes a large block of DNA
A smaller deletion can be made by cutting a plasmid at a single site. The ends of the linear DNA are then
digested with an exonuclease that removes nucleotides from both strands. The shortened piece of DNA is
then ligated to form a circle that is missing a short length of DNA about the restriction site
Designer Genes
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new proteins can be created by splicing together gene segments that encode domains not associated in nature
example includes joining a gene for an antibody with a gene for a toxin to produce a chimeric (DNA
molecule whose sequence is derived from two or more different organisms) protein that kills cells that are
recognized by the antibody. These immunotoxins are being evaluated as anticancer agents
Recombinant DNA Technology Has Opened New Vistas (Conclusion)
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Allows for mapping and the dissection of chromosomes so they can be manipulated and deciphered
Amplification of genes has allowed for sequencing
Analyses of genes and cDNA can reveal the existence of previously unknown proteins, which can be isolated
and purified
Conversely, purification of a protein can be the starting point for the isolation and cloning of its gene or
cDNA, and all one needs is a small amount of DNA or protein because of PCR
Site-specific mutagenesis opens the door to understanding how proteins fold, recognize other molecules,
catalyze reactions, and process information
Large amounts of protein can be obtained by expressing cloned genes or cDNAs in bacteria (ie insulin) or
eukaryotic cells
And the bs goes on… whatever
Okay, this covers lectures 4 and 5. Lecture 6 is not covered, but it’s only 7.1 and 7.3 in the text (5 pages). The notes
above cover the textbook and Dr. Hampson’s lecture notes completely (I hope) Oh yeah, don’t forget about the
articles Dr. Hampson handed out in class – we could very well be tested on it. Well, that’s a wrap – I hope these
notes help. Good luck with your studying – only 3 more to go!!