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Molecular Biology Exam 1


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What holds DNA strands together?
1. H-bonding (2-3 kcal/mol)
2. Hydrophobic interactions
3. Base stacking (4 kcal/mol)
adenine & guanine
cytosine, thymine, uracil
Hyperchromic shift
The increase in DNA's solution absorbance of 260nm light upon denaturation.

Sigmoidal plot
temperature at which 50% of DNA is melted.

linearally related to GC content
Molecular definition of a gene
the entire nucleic acid sequence that is necessary for the synthesis of a functional polypeptide or RNA molecule (includes promoters, terminators, etc.)
DNA sequences in concert with the proteins they help recruit. They designate where to start copying a gene. Are known as the promoter and transcription initiation site.
factors that affect melting/annealing of DNA
1. heat
2. low or high pH
3. low ionic strength
template strand (antisense)
The strand that the mRNA is formed with.
non-template strand (sense)
The strand that looks like the mRNA.
features of B DNA (4)
1. right-handed helix
2. one turn = 34A
3. distance btwn bases = 3.4A
4. helix diameter = 20A
RNA 2' OH group (2)
1. separates RNA from DNA
2. makes RNA susceptible to alkaline hydrolysis of phosphodiester linkage
functions of protein (6)
1. as enzymes
2. signal transduction
3. transport
4. vital in cell structure and shape
5. regulation of cell processes
6. maintain genetic information
codon usage
difference or distribution in codon use frequency
restriction sites that have the same sequence, but are cut at different places
factors affecting restriction enzyme cleavage frequency (4)
1. length of DNA
2. probabilistic distribution of bases in DNA
3. base composition (CG dinucleotide rare, for instance)
4. # of nucleotides in recognition sequence
Formula for determining probability of restriction sites
4^n, where "n" is the length of the recognition sequence
DNA ligase
makes a phosphodiester bond between the 5' phosphate and 3' hydroxyl
features of relaxed replication machinery
1. not under cell cycle control
2. can use stable host proteins for replication
3. can amplify plasmids by adding chloramphenicol
4. high copy
stops translation because bacterial replication stops (but plasmid replication continues)
features of stringent replication machinery (3)
1. use cellular proteins that are unstable and under cell cycle control
2. low copy
3. require a partitioning system
features of selectable markers for plasmids (2)
1. usually dominant
2. resistance to antibiotics or auxotrophic
Where are promoters found?
adjacent to the MCS
Boyer and Cohen
Performed first cloning experiment: ligated 2 plasmids together. One had tetracycline r, and the other strep r. Selected clones that had both resistances.
how to screen for recombinant molecules (pBR322)
1. Use RE to cut in amp r site (still retains tet resistance) and DNA frag.
2. Ligate frag to plasmid and transform into E. coli.
3. plate first on LB + tet, then replica plate onto LB + amp.
4. choose colonies that didn't grow on the amp plate.
Why don't restriction enzymes destroy the host cell's DNA?
Because the host cell's DNA is methylated by methylases at the restriction sites, whereas plasmid DNA is not.
restriction-modification system
the system where restriction enzymes are usually paired with methylases. Both recognize the same sequences of DNA and cleave or protect, respectively.
null allele
a gene that loses its function due to gene disruption--DNA inserted into the gene.
a tetrameric enzyme that cleaves lactose into glucose and galactose. It can also cleave x-gal to produce a blue pigment.

coded for by the lacZ gene
lacZ gene
codes for beta-galactosidase

induced by lactose or IPTG

inserted in a number of plasmids in order to screen for foreign DNA inserts.
Why is alpha complementation needed?
so that the entire lacZ gene does not have to be on an entire plasmid.
alpha complementation
C terminal is called the omega fragment and N terminal is called alpha fragment. If expressed together, the lacZ protein can still function.

Thus, omega frag can be in the host cell, and alpha on the plasmid. If a cell contains a relegated plasmid, the colony will be blue. If a cell contains a recombinant plasmid, the colony will be white because lacZ function will be lost.
plasmid copy number and size
Generally, plasmid copy number is inversely proportional to size.
What could cause a null allele?
1. out of frame insert
2. an insert that contains a stop codon
3. an insert that affects protein structure at critical points
MCS insert into lacZ'
18 nucleotides can be added in frame to the alpha frag of lacZ and a functional protein still results.
considerations for choosing a plasmid (6)
1. copy number
2. size of plasmid/insert
3. MCS
4. promoter
5. selectable marker
6. ability to screen/select
advantage of using phage vectors over plasmids
phages infect cells with greater efficiency than plasmids transform cells so yield of clones is higher with phages.

also, phages can accomodate more DNA
alkaline phosphatase
prevents vector religation by removing 5' phosphate and thus preventing ligation with itself (but can still ligate to insert DNA that contains its 5'phosphate)
filamentous phages
1. can make both ss and ds DNA (ss is packaged into phage head, ds made only during DNA replication)
2. no limit to insert size because head will grow to accomodate it
3. result in plaques on E. coli lawns
hybrids between plasmids and ssDNA phages. contain origins of replication for dsDNA plasmid and ssDNA phage.

allow high yield of ss or ds DNA.

its MCS is flanked by 2 different RNA pol promoters which allows one to isolate the ds recombinant phagemid DNA and transcribe it in vitro to produce pure RNA transcripts corresponding to either strand of the insert.
a commercial phagemid
phagemid + host alone produces ?

phagemid + phage + host produces ?

What is the use of producing ssDNA?
1. for DNA sequencing
2. to probe in a specific direction
structure of lambda vectors
contain L and R arms, containing the genes for lytic replication. The DNA insert goes in the middle.
cos sites
single-stranded cohesive ends flanking the genome of lambda phage. These sites can base pair and are ligated to form a circle.

replication takes place by a rolling circle mechanism
linking of genomes end-to-end in rolling circle replication.

genomes are separated by cos sites.
how DNA is packaged in lambda phages
lambda heads package genome equivalents (concatomers) using the cos sites to mark the beginning and end of the genome.

the heads and tails assemble to create a mature lambda particle.
cloning using charon 4 (lambda phage)
1. cut vector with EcoRI and remove middle DNA. Ligate partially digested insert DNA to L and R arms.
2. mix recombinant DNA with lambda heads and tails in vitro.
3. plate onto E. coli and collect plaques.
What insert size can the lambda phage accomodate?

Can package 78-105% of DNA (37-52kb total)
What insert size can plasmids accomodate?
less than 8kb
cosmids: definition, features, mechanism
plasmids that can be packaged into phage heads

they contain lambda cos sites and also plasmid features

because there is no lytic phage DNA, after phage infect E.coli, E. coli + cosmids result in colonies on selection plates (cosmid carries recomb DNA to E. coli)
What insert size can cosmids accomodate?
Total amount of DNA must be between 38-53kb, but since the only DNA from lambda is the cos sites, inserts of 40-50kb can be made.
circular DNA molecules that contain a replicon based on the F factor.

around 8kb in length

host needs to be deficient in homologous recombination machinery in order for the BAC to be stable.

cloning efficiency poor, DNA yield low
What insert size can BACs accomodate?
YACs: properties in yeast and bacteria, selection type, copy number
behave similarly to chromosomes in yeast (linear) and as plasmids in bacteria (circular)

contain origin of replication and selectable marker

in yeast, have sequences that function as telomeres, contain ARS, centromeres, auxotrophic markers.

selection is complementation of a prototrophy

single copy
What insert size can YACs accomodate?
~1 Mb
YACs: cloning method
1. cut at MCS to linearize and add insert DNA. Make sure one selectable marker is on one side of the insert and a second marker is on the other side.
2. select for both markers, to ensure you have L and R arms
How is a genomic library constructed?
Using overlapping fragments of DNA that have been generated by mechanical shearing or partial digestion with frequently-cutting REs.

Size select frags by gel purification
What type of vector is often used for genomic libraries from organisms with large genomes?
lambda vectors because you need fewer recombinant molecules to have confidence that you have cloned the entire genome.
What is the purpose of expression vectors?
To make the product of a cloned gene for further study.
What do you need in a good expression vector?
1. strong promoter
2. inducible promoter (drug or heat inducible)
3. ribosome binding site near an ATG codon
benefits of fusion tags
1. allow easy affinity purification (e.g, 6-His tag, GST tag, MBP tag)
2. allow visualization of protein in cell (e.g. green fluorescent tag--GFP, yellow--YFP)
using a Ni affinity column to purify proteins
1. lyse cells to release proteins
2. pour lysate through affinity column, where the Ni binds His + your protein and lets all other proteins pass through.
3. Release fusion protein with histidine or imidazole.
4. Cleave fusion protein with enterokinase
5. pass through column once more
features of a cDNA library (3)
1. contains entire protein-encoding DNA content
2. mRNA used as a starting material
3. mRNA reverse-transcribed into cDNA
how to make cDNA
1. start with mRNA and attach a poly-T primer to poly-A tail.
2. Add reverse transcriptase and dNTPs
3. Use RNase I to partially cleave RNA in the RNA/DNA hybrid, leaving string of RNA primers paired to cDNA.
4. Add DNA polymerase to find 3'-OH of RNA primers and make DNA, chewing up RNA along the way
how to make a cDNA library (already have cDNA)
1. add oligo-dC and terminal transferase to add sticky ends
2. anneal these CCC ends to a suitable vector. Transform as usual.
selection vs. screening
selection is when only the gene you want will grow (e.g. when you clone by complementation)

screening is when many things grow and you have to find your gene by a probe
how to screen plaque colonies with a DNA probe
1. nitrocellulose/nylon paper overlaid on plate phage library
2. denature with alkali solution, bake/crosslink ssDNA to filter
3. block filter with nonspecific ssDNA to lower background noise, hybridize ssDNA to labeled probe
4. detect by autoradiography
how to screen colonies with an antibody probe
1. blot proteins from phage plaques to a filter.
2. incubate filter with an antibody directed against the protein of interest
3. filter is then incubated with labeled Staph protein A, which binds to most antibodies, and thus will only bind where your protein has bound antibodies
4. detect by autoradiography
where do antibody probes come from?
purified protein or from a homologous protein
where do DNA probes come from?
1. from a homologous gene (lower stringency)
2. can predict the sequence within the coding region of a highly conserved gene and construct synthetically
considerations for designing an oligonucleotide probe
1. the longer, the better. (usually over 14 bp)

2. are degenerate. Know the number of oligos you have to use by multiplying the number of codons that code for each amino acid.
PCR, briefly
1. denature at 95 degrees
2. anneal primers at 50 degrees
3. elongate strandfs at 72 degrees.
4. repeat 20-35 times.
What is RACE?
rapid amplification of cDNA ends

a method to fill in the missing pieces of cDNA.
RACE: method
1. hybridize incomplete cDNA to mRNA and use reverse transcriptase to extend cDNA to 5' end of mRNA.
2. Use terminal transferase and dCTP to add C residues to 3' end of cDNA and use RNase H to degrade mRNA.
3. Use oligo(dG) primer and DNA polymerase to synthesize a second strand of cDNA
4. perform PCR with oligo(dG) as forward primer and and oligonucleotide that hybridizes to the 3' end of the cDNA as reverse primer.
Choosing a library:
cDNA or genomic?
cDNA: if you want to clone protein-encoding genes and use them for protein expression

genomic DNA: if you want the DNA sequence, to look at promoter regions, see introns
Choosing a library:
which vector system?
Consider size of inserts, whether we want to be able to generate proteins directly, ss or dsDNA
Choosing a library:
which host?
E. coli or S. cerevisiae (if eukaryotic equipment needed for protein modification)
sodium dodecyl sulfate: a detergent (denatures proteins) that coats proteins with negative charges, which serves to make then migrate to the anode and masks original charge so they migrate by size.
used to separate proteins by mass
2-D gel electrophoresis
1. isoelectric focusing: protein mixture is electrophoresed and move until they reach their isoelectric points.
2. gel is removed from tube and run on an SDS-PAGE gel and separate according to size

allows greater resolution of proteins
ion-exchange chromatography
separates proteins based on charge.
1. run proteins through column with charged resin (DEAE-Sephadex)
2. elute with a buffer of increasing ionic strength and collect fractions.
gel-filtration chromatography
run proteins through a column filled with porous resin that let smaller substances through, but not larger ones. larger substances flow through column more quickly.
radioactive emissions expose film, which when developed, produces dark spots
more accurate in quantifying amount of radioactivity than autoradiography.

fix a hybridized blot to a phosphorimager plate, which absorbs beta rays. The molecules on the plate will remain excited until a phosphorimager scans the plate with a laser. The beta-ray energy is then detected by a computer.
liquid scintillation counting
uses radioactive emissions from a sample (inside scintillation fluid) to create photons of visible light that a photomultiplier tube can detect.
Southern blotting
labeled DNA (or RNA) probes are hybridized to DNAs of similar sequences.
1. DNA run on a gel is denatured with an alkaline buffer and transfered (ssDNA) to a nitrocellulose filter by diffusion or electrophoresis.
2. hybridize blot to labeled DNA or RNA prbe and detect labeled bands by autoradiography.
DNA fingerprinting
uses Southern blotting and DNA probes to detect variable sites (minisatellites)
DNA typing
probes hybridize to a single DNA locus that varies from one individual to another (instead of to many loci as in a fingerprint)

extremely sensitive
Northern blotting
use to measure gene activity

electrophoretically-separated RNA is on the blot and is hybridized to a labeled cDNA probe.

the intensity of the bands reveal relative amounts of specific RNA in each

position of bands indicate length of RNA
in situ hybridization
identifies which chromosome a given gene is on

partially denatures DNA in chromosomes to create ss regions that can hybridize to a labeled probe.

FISH: probe is fluorescently labeled
Sanger Chain-Termination method of DNA sequencing
1. a primer is hybridized to ssDNA and mixed with the Klenow frag of DNA polymerase and dNTPs. One type of dideoxy NTP is included to terminate replication after certain bases. 4 separate rxns.
2. Electrophorese products. Read gel from bottom up. This is the sequence of the non-template (sense) strand.
site-directed mutagenesis
1. Denature a methylated plasmid with heat and anneal mutagenic primers.
2. Perform about 8 rounds of PCR to amplify and a Pfu (stable) polymerase.
3. Treat DNA from PCR with DpnI, which cuts at methylated sites, and thus gets rid of all wild type DNA.
4. Transform into E. coli. (This is an example of selection)
Why is S1 mapping used?
used to locate 5' or 3' ends of RNAs and to quantify amount of RNA in a cell at a given time
S1 mapping: method
1. cut cloned DNA with REs and remove unlabeled phosphates.
2. Label 5' ends with gamma-P ATP and cut with SalI, so that just one end is labeled.
3. denature and hybridize with RNA transcript.
4. treat hybrid with S1 nuclease, which digests ss DNA and RNA.
5. denature hybrid and electrophorese to see how large it is.
6. length of protected probe indicates position of transcription start site.
primer extension
can map 5' end of RNA transcript to the nucleotide

1. harvest cellular RNA and hybridize a labeled oligo DNA probe to transcript
2. use reverse transcriptase to extend primer by synthesizing complementary DNA
3. denature the hybrid and electrophorese the labeled primer.

This can indicate the transcription start-site to the base.
run-off transcription
1. cut cloned gene with RE and transcribe in vitro. When RNA pol reaches end of shortened gene, it falls off.
2. the size of the run-off transcript can be measured by electrophoresis and corresponds to the distance between the transcription start site and the restriction site at the 3' end.
G-less cassette
a promoter is fused to a dsDNA cassette lacking G's in the non-template strand

1. transcribe a template with a G-less cassette downstream of the promoter in vitro in the absence of GTP.
2. Electrophorese the labeled transcript and autoradiograph the gel.

The intensity of the signal indicates how actively the cassette was transcribed.
nuclear run-on transcription
1. isolate nuclei in the process of transcription and incubate with nucleotides so that transcription can continue. Include 1 type of labeled nucleotide in reaction.
2. do a dot blot assay, where ssDNA from the transcribed genes is blotted to nitrocellulose and hybridized to the labeled run-on transcript.

The more active the transcription of the gene, the more intense the labeling.
reporter gene
a gene attached to a promoter or translation start site, and used to measure the activity of the resulting transcription or translation. The reporter gene serves as an easily assayed surrogate for the gene it replaces.

e.g. lacZ, cat
Western blotting (immunoblotting)
proteins are electrophoresed, blotted to a membrane, and probing with a specific antibody or antiserum to detect a particular polypeptide.
a technique in which labeled proteins are reacted with a specific antibody and then precipitated by centrifugation. The precipitated proteins are detected by electrophoresis and autoradiography.
nitrocellulose filter binding assay
label dsDNA, mix it with a protein and pass it through a nitrocellulose filter. Then meausre the amount of radioactivity on the filter.
gel mobility shift assay (EMSA)
since small DNA has a much higher mobility than DNA bound to a protein, one can label a short DNA fragment, mix it with protein, and electrophorese it. Detect with autoradiography.
DNase footprinting
uses DNase to cleave DNA. Can determine where protein is bound by looking at the results of electrophoresis and finding a gap in fragments (this area is protected by the protein)
DMS footprinting
Can determine where a protein is bound more accurately than with DNase, which is a macromolecule. DMS methylates DNA, and these sites can be cleaved by the Maxam-Gilbert sequencing reagant. Visualize by electrophoresis/autoradiography to see where protein is bound.

Sometimes, bands representing where the protein is bound grow darker as protein concentration is increased, which is because sometimes DNA is unwound at the binding site, making it more vulnerable to methylation.
footprinting with hydroxyl radicals
organometallic complexes containing copper or iron generate hydroxyl radicals that break DNA, except where bound to protein.

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