Appendix: some lab techniques (need progress)

Enzyme-linked immuno-sorbet assay [ELISA] (page 189)

As the name suggests, ELISA is a biochemical technique that utilizes antigen-antibody interactions (‘immuno-sorbency”) to determine the presence of either

1. antigens (like proteins or cytokines) or

2. antibodies

in a sample, (such as cells recovered from a tumor biopsy or a patient’s serum).

Figure 1 illustrates the basic protocol when testing for the presence of a specific antigen.

1. The experimental wells are coated with antibodies that are specific for the target antigen.

2. A sample of serum or cell extract is added to the wells.

3. The antibodies immobilize the antigen by binding to it (if the antigen is present in the sample)

4. Any unbound proteins remaining in the sample are washed away

5. An enzyme-linked antibody that also recognizes the target protein is added to the wells.

6. The wells are filled with a solution that changes color in the presence of the detection enzyme (the one with the antibody linked to it in step 5). A color change indicates the target protein was present in the sample; no color change means the protein was absent.

Figure 2 (when testing for a specific antibody)

When testing for the presence of a specific antibody in a sample,

1. the antigen (for which the antibody is specific) is first allowed to adhere directly to the wells.

2. The sample is added as above,

3. and then mixed with enzyme-linked antibodies (see figure 2).

Radioimmunoassay (RIA)

A radioimmunoassay (RIA) is a sensitive in vitro laboratory technique used to measure the concentration of tiny amounts of an antigen (such as hormones, drugs, or proteins) in a biological sample using antibodies and radioactive isotopes.

RIAs are similar to ELISAs but use radio-labeled antibodies rather than enzyme-linked antibodies. Thus, the presence of target proteins or antibodies is assayed by measuring the amount of radioactivity instead of a color change. RIAs are more extensively used in the medical field to measure the relative amounts of hormones in patient’s sera (see figure 3).

-RIA is used to measure the relative amounts of hormones in patient’s sera.

1. a known amount of radiolabeled antigen (for example, insulin that was synthesized with ¹²⁵I-labeled tyrosines) is incubated with a known amount of antibody that is specific to the antigen.

2. The insulin:antibody complexes are isolated

3. The total amount of radioactivity is measured

4. Unlabeled insulin (also called cold insulin) is mixed into the solution in increasing amounts. The cold insulin competes with the labeled insulin (hot insulin) for the antibody.

5. As more cold insulin is added, less total radioactivity is recovered and measured. This competition assay helps formulate a standard curve (see Figure 4)

6. Steps 1-3 are repeated using patient serum instead of cold insulin. The standard curve is used to extrapolate the amount of insulin that is circulating in a patient’s serum.

Electrophoresis

• electro-: electron

• -phoresis: carrier

The purpose of electrophoresis is to separate things by size (for example, nucleic acids or protein) or by charge (for example, proteins or individual amino acids).

1. A “gel” is made out of either acrylamide or agarose by solubilizing the acrylamide or agarose, pouring it into a rectangular mold, and then allowing it to cool and solidify. Acrylamide and agarose form “nets” as they solidify; the more acrylamide or agarose used in the initial solution, the smaller the pores of the nets.

2. The mold used to pour the gel creates wells in the gel into which samples can be loaded.

3. An electrical current is applied such that the end of the gel with the wells is negatively charged and the opposite end is positively charged.

4. This causes the samples to migrate toward the positive pole, according to size, smaller things migrate faster (because they fit more easily through the pores of the gel) and larger things migrate more slowly.

For example:

1. Isolate the sample DNA from cells

2. Expose the DNA to enzymes called restriction endonucleases (see section A.5), which cleave the strands of DNA into smaller fragments of varying size. This may not be necessary in some cases

Restriction endonucleases are specialized proteins produced by bacteria and archaea that cut double-stranded DNA at specific nucleotide sequences called restriction sites

3. Add a loading dye to the DNA sample. This makes the sample visible as it is being loaded into the gel. Loading dye also contains a chemical to help inhibit DNA degradation. Finally, glycerol in the loading dye makes the sample more dense than in the surrounding buffer, which means the DNA sample sinks to the bottom of the gel wells.

4. Load the mixture of fragments into the gel wells and apply the electrical current (this is called “running a gel”).

5. Each strand of DNA (remember DNA is negatively charged!) migrates toward the positive end of the gel, but the smaller fragments migrate more quickly, so they are found farther from the wells at any point in the experiment. You run the samples alongside a “standard” lane, which contains fragments of known size (this helps identify the size of the unknowns).

6. Visualize the bands of DNA in the gel. This is done using a dye that binds to nucleic acids and fluoresces when exposed to UV light. This dye is typically added to the gel when it is being made, but it can also be applied after the gel is run. The size of each DNA band can be approximated by comparing it to the ladder.

take a picture of figure 5 and put it here.

In addition to determining their sizes, fragments of DNA (or RNA) in an electrophoresed gel can be transferred to a more solid and stable membrane in a process called “blotting”. There are several types of blots used in biology laboratories.

Blotting

Blotting is the transfer of DNA or proteins from an electrophoresis gel to a nitrocellulose of PVDF membrane.

Once transferred, further experiments can be run to isolate or detect a particular nucleic acid fragment or protein (called “probing”). Blotting is classified by the type of molecule being probed.

Southern Blotting

Southern blotting allows you to detect the presence of specific sequences in heterogenous sample of DNA. This process also allows you to isolate and purify target sequences of DNA for further study.

1. Separate the DNA fragments on a electrophoresis gel (electrophoresis)

2. Transfer the fragments to a nitrocellulose membrane (southern blotting)

3. The filter is “probed” for their target DNA sequence. Hybridization probes are short single-stranded sequences of nucleic acid (usually DNA) that have two important features:

-they are complementary to (and thus will base-pair with) a portion of the target DNA sequence.

-they are constructed with radiolabeled nucleotides, which allows the visualization of the target sequence with special film.

Probes are often engineered to complement mutations or certain gene rearrangements, making Southern blotting a useful diagnostic tool.

put figure 6 here

Northern Blotting

Northern blotting is for RNA.

Northern blotting is almost identical to southern blotting, except that RNA is separated via gel electrophoresis instead of DNA.

The rest of the process is the same; once the RNA has been separated on a gel, it is transferred to a nitrocellulose membrane and detected via radiolabeled nucleic acid probe. This technique allows you to determine whether specific gene products (normal or pathologic) are being expressed (if their mRNA is present in a cell, they are probably being translated to a protein).

Western Blotting

Western blotting allows you to detect the presence of certain proteins (and the size) within a sample and also serves as as diagnostic tool. You are able to determine, for example, whether cancer cells express certain tumor-promoting growth receptors on their surface.

1. Cells are collected and solubilized in detergent to release their cytoplasmic contents.

2. Cell lysates, which contain hundreds of different proteins, are denatured (meaning they lose their secondary and tertiary structures).

3. Lysates and a ladder are loaded onto a gel. Similar to nucleic acid gel electrophoresis, a ladder is used so protein size can be compared to a standard.

4. An electric current is applied. Because of the detergent used, the proteins are all negatively charged.

5. Therefore, they migrate toward the positive electrode, with the smaller proteins migrating the farthest from the wells.

6. The separated proteins from the gel are transferred to a nitrocellulose or PVDF membrane

7. The membrane is probed for the target protein. Probing for proteins in Western blotting differs from probing in Southern or Northern blotting in that antibodies are used as the probes rather than nucleic acids. This is similar to the technique in ELISA; a primary antibody is used first, which will recognize only the target protein via its antigen-binding portion. Then, an enzyme-linked secondary antibody is used that recognizes the constant region of the primary antibody. The enzyme on the secondary antibody will fluoresce when a detection substrate is added, and this light can be photographed with special film. The target protein will show up as a band with an intensity that is proportional to the abundance of the protein in the sample (see Figure 7)

Eastern Blotting

Several variations of eastern blotting have been reported, but these tests are not commonly used in molecular biology labs. Eastern blots are used to analyze post-translational modification of peptides, such as the addition of lipids or carbohydrates. The details of this protocol depend on the specifics of the experiment.

Recombinant DNA

Recombinant DNA (rDNA) is an artificially created DNA molecule formed by joining genetic material from two or more different sources or species, producing sequences not naturally found together in a genome.

In the past twenty years, a major change has occurred in biology that has allowed it to not only describe the mechanisms of life, but also manipulate living organisms. The cloning and sequencing of genes, production of recombinant DNA, and the subsequent production of recombinant proteins for use as therapeutic agents in medicine have now become commonplace procedures.

A recombinants protein is one which has been obtained by transcribing and translating a novel combination of DNA (recombinant DNA) from different organisms.

1. recombinant DNA

2. recombinant protein obtained

For example,

1. the gene for human insulin can be placed in a bacterial plasmid (described below).

2. Bacteria with the plasmid will then produce insulin that can be used to treat diabetes. To a large extent, these advances are due to the development of new technologies for the handling of DNA, such as the discovery of restriction endonucleases that cleave particular DNA sequences.

Restriction endonucleases are bacterial enzymes that recognize specific sequences of DNA and cut the double-stranded molecule in two peices.

Restriction:• Derived from the observation in the early 1950s that certain bacteria could restrict or limit the growth and replication of invading bacteriophages (viruses that infect bacteria).

Endo-:• "within" or "inside".

nuclease:• A combination of nucleic (referring to nucleic acids like DNA or RNA) and the suffix -ase, which is the standard chemical designation for an enzyme that catalyzes a reaction.

A nuclease is an enzyme that cuts nucleic acids. An endonuclease cuts the middle of a DNA chain (contrast with exonucleases, which nibble nucleotides from the ends of DNA chains). They are isolated from bacteria and used in the lab. Their natural role in the bacterium is to destroy viral DNA that gets injected into the cell; thus, they restrict the reproduction of hostile viruses.

Restriction enzymes have found great use in molecular biology, where they have permitted manipulation of genes to create recombinant DNA. For example, in Figure 8 below, the cutting-specificity of a restriction enzyme known as EcoRI is shown (other restriction enzymes cut at different sequences). The free ends of the DNA molecule that were complementary are known as sticky ends since they are able to base pair with other DNA molecules with similar sequences.

Put figure 8 here

Study the sequence shown in Figure 8. Notice anything in particular?

1. If you read the top strand from left to right (5’ to 3’), it begins with GAATTC.

2. Now read the bottom strand from right to left (still 5’ to 3’), but only read the six nucleotides on the left side of the chain. It says GAATTC (same as above)!

3. Just looking at these six nucleotides, we see that the chain possess two-fold rotational symmetry. The six 5’ nucleotides of the top chain are the same as the six 3’ nucleotides of the bottom one. Sequences with two-fold rotational symmetery are known as palindromes. Many restriction enzymes recognize palindromic sequences.

palindrome is a word, number, phrase, or sequence that reads the same forwards as it does backwards

Two-fold rotational symmetry (also called a half-turn symmetry or C₂ symmetry) means an object looks completely identical after being rotated by 180 degrees around a central point or axis. Because a full circle is 360 degrees, the shape repeats its appearance twice during a complete 360-degree rotation (360° ÷ 180° = 2)

1. When a fragment of double-stranded DNA is created by cutting with a restriction endonuclease,

2. the fragment can be inserted into DNA from any source that was also digested by the same restriction endonuclease.

-the restriction endonucleases need to match, why? because they recognize and cut the exact same specific DNA sequence (the recognition site).

For example,

1. EcoRI-generated DNA fragments from a human can be isolated,

2. mixed with EcoRI-digested DNA from a bacterial plasmid,

3. then joined by the enzyme DNA ligase.

4. Hybrid DNA produced in this fashion is referred to as recombinant DNA.

recombinant DNA= hybrid DNA

Figure 9 (uses the same restriction endonuclease to produce the recombinant DNA)

EcoRI is a restriction endonuclease

Some restriction enzymes generate DNA with blunt ends rather than sticky ends. That is, the 3’ and 5’ ends at the cut site are even, with no overhanging bases. Ligating blunt ends together is less specific, and restriction sites may or may not be retained.

-For example, if the same blunt-cutting restriction enzyme is used on both pieces of DNA, the restriction site will be maintained after ligation.

-If different blunt cutting enzymes are used, the products can be ligated together, but neither restriction site will be maintained.

In Figure 10, a bacterial plasmid was digested with the restriction enzyme SmaI (which was a blunt cutter and recognizes the restriction site CCCGGG.) A human gene of interest was digested with the restriction enzyme. EcoRV (which is a blunt cutter and recognizes the sequence GATATC.) Notice that both these restriction sites are six base pair palindromes. Because both enzymes generate blunt ends, these products can be ligated together. However, the recombinant DNA has lost the restriction sites for both enzymes. The DNA that remains is a combination of the two blunt sites (CCCATC and GATGGG), and it cannot be digested with either SmaI or EcoRV.

Figure 10 (hybrid DNA produced using different restriction endonucleases resulting in blunt ends)

• Same restriction enzyme (EcoRI): Produces matching sticky ends, allowing the foreign DNA and plasmid DNA to pair easily and form recombinant DNA.

• Different restriction enzymes (EcoRI and SmaI): Produce incompatible ends (sticky and blunt), so they cannot directly pair through complementary base pairing (they can be joined by being made compatible through a technique called linker addition or adaptor ligation to make incompatible DNA ends compatible)

Plasmids

Plasmids are small circular ds-DNA molecules found in bacteria that are capable of autonomous replication (replication that is independent of chromosome replication).

A plasmid is a small, circular, double-stranded DNA molecule that is separate from a cell's chromosomal DNA and can replicate on its own. While not essential for normal daily survival, plasmids give host organisms useful traits to handle stressful or changing environments.

Natural Functions in Bacteria

• Antibiotic Resistance: Carry resistance genes (R-plasmids) that protect bacteria from toxic drugs and allow the trait to spread.

• Horizontal Gene Transfer: Use specialized transfer genes (F-plasmids) to build a connection (pilus) and share genetic material with other bacteria via conjugation.

• Virulence and Toxins: Increase a pathogen's ability to cause disease or produce competitive toxins (Col-plasmids) that kill rival bacteria.

• Bioremediation and Metabolism: Enable bacteria to digest hard-to-break-down pollutants, hydrocarbons, or toxins in the environment (degradative plasmids)

Plasmid replication still requires an origin of replication (ORI), which affects the copy number of the plasmid.

-Some plasmids have strong ORIs, leading to hundreds of plasmid copies per cell.

-Other ORIs are less efficient, leading to only a few copies of plasmid per cell.

In addition, plasmid segregation during binary fission is not regulated.

-For high copy plasmids, both daughter cells will most likely end up with copies of the plasmid.

-For lower copy plasmids, one daughter cell could get all copies of the plasmid, while the other daughter cell gets none. Plasmids used in laboratories are almost always high copy plasmids. The presence of a large number of copies is convenient, since it allows for isolation of plasmid DNA with identical sequences.

Plasmids have been manipulated by recombinant techniques to propogate and express foreign genes in bacteria. In addition to an ORI, they also contain a multiple cloning site, which has restriction sites for dozens of restriction enzymes. This means the plasmid can be digested and any desired sequence with complementary ends can be ligated into the plasmid.

Furthermore, plasmids have a drug resistance gene that helps select and isolate bacteria possessing the plasmid from other bacteria.

For example,

1. bacteria containing a plasmid with the ampicillin-resistance gene are able to grow in the presence of the antibiotic ampicillin (and are AmpR, or resistant), while bacteria that do not possess the plasmid will die in the presence of ampillin, only those bacteria that possess and express the plasmid can grow and maintain colonies (see Figure 11). Tetracycline, penicillin, and streptomycin are other commonly used prokaryotic selection agents.

Put figure 11 here

Bacterial expression plasmids need extra components. A prokaryotic promoter and start site allow expression of an inserted gene in the bacterial host. The promoter can be either constitutively active or inducible upon addition of a chemical.

Bacterial transformation

Bacterial transformation is the process where a bacterial cell takes up foreign genetic material (usually a plasmid or DNA fragment) from its environment and incorporates it, gaining new traits.

Plasmids can be easily reintroduced into bacterial cells via transformation. Transformation is a naturally occurring process, but only a very small percentage of bacteria are naturally willing to accept pieces of DNA floating around in their environment. These “competent” bacteria express special machinery that translocates the hydrophilic DNA across the lipid membrane.

More often, the bacteria (or other cell types) must be coaxed to take up the plasmid. There are several ways to do this; the cells maybe cooled in calcium chloride, and then heart shocked to facilitate plasmid uptake. In another (called electroporation), an electric field is applied and this pokes holes in the membrane, which allows the plasmid to diffuse into the cell.

1. Once inside the bacteria,

2. the plasmid will be exposed to the host’s replication machinery, which replicates the plasmid (remember, it has its own origin of replication). If you are working with an expression plasmid, the DNA is also exposed to bacterial transcription machinery (remember the plasmid contains the proper promoters and start signals). Newly synthesized mRNA can then access host ribosomes, which translate the encoded protein; on completion of translation, the cells are lysed to release the protein.

If the plasmid remains with the cytosol of the bacterium, the transformation is referred to as transient. In contrast, some plasmids are constructed to integrate within the host’s genome. These stable transformations allow the plasmid to be replicated each time the bacterium replications its own genome.

Complementary DNA

Complementary DNA (cDNA) is synthetic DNA that is reverse-transcribed from messenger RNA (mRNA) or microRNA template using the enzyme reverse transcriptase. .

Complementary DNA (commonly abbreviated as cDNA) gets its name because its nucleotide sequence is a direct base-pairing complement to a messenger RNA (mRNA) template from which it is synthesized.

Many applications of DNA technology involve expressing eukaryotic genes in prokaryotic cells such as E.colo. This is conceptually, simple: all you have to do is get eukaryotic DNA into a plasmid and get the plasmid into a bacterium, and the bacterium should express the gene. However, there are two main problems that need to to be overcome.

1. prokaryotes lack the equipment neccessary for splicing out introns.

2. many eukaryotic genes are extremely long, making them hard to work with.

One way to overcome these obstacles is to work with eukaryotic complementary DNA. cDNA carries the coding sequence for genes, lack introns, and thus are shorter.

1. Complementary DNA (or cDNA) is produced from fully spliced eukaryotic mRNA. This is accomplished by reverse transcriptase from retroviruses. Reverse transcriptase reads an RNA template and builds complementary DNA.

2. Therefore, RNA can be isolated from a eukaryotic cell and converted into cDNA by the addition of reverse transcriptase, some generic primers and dNTP building blocks in buffer. cDNAs carry the complete coding sequences for genes, but they lack introns (and thus are smaller than the genomic sequence of the gene).

3. Once a cDNA is ligated into a bacterial plasmid and bacteria are transformed with this plasmid, they can produce the protein encoded by the cDNA.

cDNA libraries are also commonly generated. This is where each of the thousands of mRNAs being generated by a given cell type or tissue are converted into cDNAs. Each cDNA is then cloned into a plasmid. This generates thousands of plasmids (the library), with each one containing one cDNA molecule. cDNA libraries can be compared across tissue types of a certain organism (brain verses liver, for example) to study tissue-specific gene expression.

Artificial Chromosomes

Plasmids can only carry inserts up to a certain size. If large inserts are required, artificial chromosomes can be used.

1. Bacterial artificial chromosomes (BACs) typically carry inserts of 100 to 350 kilobase pairs (kb),

2. while yeast aritifical chromosomes (YACs) can carry inserts between 100 and 3,000 kb. In other words, BACs can easily carry up to 350,000 base pairs of DNA, and YACs can contain up to 3 million base pair inerts.

BACs

BACs contains sequences to allow replication and regulation of copy number, partition genes that promote their even distribution after bacterial cell division, and a selectable marker for antibiotic resistance. BACs that express inerted sequences also contain promoter regions.

BACs can be used to study inherited diseases that involve complex genes with several regulatory sequences and promoters upstream of the coding sequence. The entire gene can be cloned into a BAC, and this can be used to model genetic diseases in mice. For example, both Alzheimer’s disease and Down syndrome as herpesviruses, poxviruses, and coronaviruses. The infective BACs initiate viral infection in the host cell and have facilitated research on these viruses.

YACs

YACs contain a centromere, telomeres, and sequences that function as replication origins. They also typically contain a gene that allows tryptophan and pyrimidine biosynthesis, allowing for selection of auxotrophic cells that contain the artificial chromosome. This system works similarly to antibiotic selection in bacteria.

YACs are less stable than BACs, but YACs can be used to study proteins that require post-translational modification (because yeasts are eukaryotes).

Both BACs and YACs were initially used in the Human Genome Project, to help make chromosome maps. However, YACs were eventally abandoned because they are less stable than BACs. Despite this, they do have one major advantage: because yeast cells are eukaryotic, YACs can be used to express and study proteins that require post-translational modification.

Eukaryotic Plasmids

Eukaryotic plasmids also exist. They require many of the same components as bacterial plasmids. Eukaryotes use different selection agents, usually either puromycin or neomycin. They also require different promoters in expression plasmids, as well as a poly-adenation signal downstream of the inserted gene, to terminate transcription.

Uromycin and neomycin (G418) are two of the most widely used selection agents for eukaryotic cells. They function by killing untransfected cells, leaving behind only the cells that successfully integrated the desired resistance gene.

Eukaryotic plasmids can be introduced into mammalian host cells via transfection (Transfection is the process of artificially introducing nucleic acids—such as DNA, mRNA, siRNA, or CRISPR components—into eukaryotic (animal) cells.).

Similar to transformation, there are several experimental options for transfection.

1. Cells can be chemically transfected, usually using caclium phosphate precipitates or plasmid packaging in liposomes. These lipid vesicles mask the plasmid, but they deliver it to the interior of the cell by fusing with the plasma membrane. Non-chemical options for transfection include electroporation, optical transfection with lasers, or shooting the DNA coupled to a gold nanoparticle into a cell nucleus using a gene gun.

Viruses can also deliver DNA into eukaryotic cells, a process called viral transduction. Transduced cells can express genes carried by the viral vector.

Transformation: bacteria gets DNA from the environment

Transfection: artificially introducting nucleic acids into eukaryotic animals

Viral Transduction: viruses deliver DNA into eukaryotic cells

Polymerase Chain Reaction

Polymerase Chain Reaction (PCR) is a quick and inexpensive method for detecting and amplifying specific DNA sequences, screening hereditary and infectious diseases, and fingerprinting DNA. Designed to generate myriads copies of single template sequences, PCR allows the amplification and subsequent analysis of very small samples of DNA.

Let’s say that PCR is to be used to determine whether a certain viral gene has been integrated within a bacterial host genome.

1. A nuclear extract of the bacteria is obtained.

2. Then primers are carefully constructed that will help locate the viral gene (if it is present within the host). Primers are engineered DNA oligonucleotides (~15 bases of single stranded DNA) that will recognize and base pair with specific DNA sequences; in this example, the primers will each recognize a 15-base stretch of the viral gene. Two primers, which will flank a total of ~10 kb of DNA, are used.

3. The “forward primer” will recognize a 15-base stretch at the 3’ end of the antisense strand,

4. The “reverse primer” will recognize a 15-base stretch at the 3’end of the sense strand.

5. When base paired to their respective gene sequences, the primers will bookend (on opposite sides) the intervening target gene segment (see Figure 12).

the antisense strand acts as the direct template for making messenger RNA (mRNA), while the sense strand matches the resulting mRNA sequence.

Figure 12 here

The primers have free 3’hydroxyl groups to which dNTPs can be added in a 5’ to 3’ direction. This will allow for elongation of complementary strands of DNA. The bacterial DNA is mixed with multiple copies of the forward and reverse primer, lots of dNTP bases, a heat-sensitive DNA polymerase, and ions into a buffer. The mixture is then placed into a PCR machine, which will carry out these three basic steps (see Figure 13):

Step 1: Initialization: The sample is heated to ~95 C°. Heating the sample “melts” the hydrogen bonds that hold the ds-DNA together, which creates single-stranded DNA.

Step 2: Annealing: The sample is cooled to ~55 C°. At this temperature, the primers base-pair with the template strands

Step 3: Elongation: The sample is heated to ~ 72 C°. Using the primers as starting points, the heat-sensitive DNA polymerase (usually Taq polymerase isolated from algae that thrive in hot springs) elongates strands of DNA that are complementary to each of the template strands. Each strand is polymerized in the 5’ to 3’ direction. Any mismatched primers will dissociate from the template strands and will not be extended (this helps ensure the purity of the PCR product). Longer DNA targets take longer to synthesize, so the length of the elongation step depends on the length of the product DNA.

Insert Figure 13 here

Each of the cycle of three steps takes between 0.5 and 5 minutes, depending on the length of the target DNA product (and subsequent length of the elongation step). Because two new complementary strands are synthesized for each template strand in the sample, the PCR product grows at an exponential rate, yielding over a billion copies in just 30 cycles. The sample of DNA is separated via electrophoresis and stained to visualize the products, including the amplified viral gene segment (if present).

Reverse Transcriptase Polymerase Chain Reaction (RT-PCR)

Reverse transcription polymerase chain reaction (RT-PCR) is a laboratory technique that converts RNA into complementary DNA (cDNA) and then amplifies specific target DNA sequences using Polymerase Chain Reaction (PCR).

This is an extension of classic PCR and is used to detect the relative expression of specific gene products.

RT-PCR provides a gauge of gene transcription by measuring the amount of target mRNAs.

To conduct an RT-PCR experiment, all of the mRNAs from within a cell population are:

1. isolated

2. converted to complementary DNA (cDNA) using the enzyme reverse transcriptase

3. This library of cDNAs is then subjected to PCR, using primers specific for a certain gene of interest.

4. If the gene was actively transcribed at the time of harvest, its mRNA will have yielded a cDNA, which will be amplified by the PCR reaction and visualized on a gel.

Quantitative Polymerase Chain Reaction (qPCR)

Quantitative PCR (qPCR), also known as real-time PCR, is a molecular biology technique that amplifies and simultaneously measures the amount of a targeted DNA molecule in real time.

In quantitative PCR (qPCR, also called real time PCR), the PCR product is both detected and quantified, either as an aboslute number of copies or as a relative amount normalized to control.

The amplified DNA is detected in “real time'“, as the reaction progresses.

The detection process can either use a dye that is fluorescent and binds DNA or a fluorescent oligonucleotide probe which hybridizes to the sequence of interest.

qPCR can be performed on either DNA or cDNA templates, meaning it can give information on the presence and abundance of a particular DNA sequence in samples (if DNA is template) or on gene expression (if the template is cDNA).

DNA Sequencing and Genomics

DNA sequencing is a method by which scientists can determine gene sequences.

The most widely used DNA Sequencing method (the Sanger Technique) hinges on a simple yet important structural characteristic of DNA molecules. The ringed ribose of a dNTP has various substituents attached to its carbons: a nitrogenous base at the 1’ carbon, a hydrogen at the 2’ carbon (recall that a hydroxyl group occupies this site in RNA), a hydroxyl group at the 3’ carbon, and a string of three phosphates at the 5’carbon. The 3’ carbon hydroxyl group serves as the binding site for another dNTP. Without a free 3’ carbon hydroxyl group, dNTPs could not be linked together, and DNA synthesis would not be possible.

The Sanger Technique utilizes a modified dNTP, which lacks the 3’carbon hydroxyl group. These dideonucleotide triphosphates (ddNTPs) maintain their 5’ carbon triphosphate moiety and can be incorporated normally into a growing DNA molecule, however, because they are lacking the 3’ carbon hydroxyl group, no further bases can be added to them. Thus, these ddNTPs terminate strand elongation at the point of their insertion. The basic protocol is as follows (Figure 14):

step 1: obtain a sample of DNA to sequence

step 2: denature the DNA into single strands

step 3: Mix the sample of DNA with radiolabeled primers, DNA polymerase, and a mixture of dCTP, dTTP, and dGTP, dATP, and ddATP (with the dideoxy form making up 1 percent of the adenine base population). This step of the assay will yield a population of newly synthesized DNA fragments, varying in length, each complimentary to the template strand and covalently bonded to a radiolableed primer at the 5’ end (this will aid in the detection of the newly synthesized fragments later). The variety in length of the fragments results from the random insertion of a ddATP into the growing chain.

step 4: Conduct three more seperate reactions as in the previous step, using each of the three other bases in dideoxy form (ddCTP, ddGTP, and ddTTP)

step 5: seperate the fragments via gel electrophoresis, running each reaction from Steps 3 and 4 in a separate lane

step 6: transfer the fragments to a membrane and visualize them in radio-sensitive film

Put figure 14 here

The smallest fragment (i.e., the fragment that migrates the farthest from the well) is a primer with only a single ddNTP attached to it. The lane is ran in corresponds to the first base incorporated into the strand which is the first base of the sequence of the complimentary strand. The second smallest fragment is a primer with two bases attached; this fragment ran in the lane corresponding to the base at the second position in the complimentary strand (see Figure 15). Reading the membrane from bottom (farthest from the wells) to top (closest to the wells) indicates the sequence (in the 5’ to 3’ direction) of the complimentary strand. Remembering the simple rules of base-paring (A:T and C:G), you can easily extrapolate the sequence of the template strand.

Put figure 15 here and decipher it.

Genomics

Genomics studies the genome—complete set of DNA.

The genome of bacterium Haemophilis influenzae was the first to be sequenced and published in 1995. Since them, the genomes of hundresds of organisms have been sequenced and published, including humans and model organisms commonly used in biology laboratoies (e.g., E.coli, D. melangaster, etc.) Researchers and clinicians have recently started sequencing the genome of many cancers, which allows comparitive studies between different types and subtypes of cancer, as well as a better understanding of how the cancer is different from a normal one.

Genomic Sequencing (genetic linkage map)

Genomic Sequencing is done in two ways, which can be complementary.

1. Genetic linkage map, with several hundred markers per chromosome. This map is then refined to a physical map by preparing YAC or BAC libraries containing large chromosomal fragments. The library is put in order, then gradually cloned into libraries containing smaller and smaller fragments. Each of these small fragments are eventually sequencd and assembled into an overall sequence.

genetic linkage map (or genetic map) is a diagram that shows the relative positions of genes or genetic markers along a chromosome. It is built by measuring how often different traits or markers are inherited together during cell division (meiosis).

2. Whole-genome shotgun approach, where chromosomes are cut into small fragments, which are cloned and sequenced. This strategy skips generating maps, and because of this, requires much more extensive analysis of sequencing data by computers in order to align fragments.

Genomic data can lead to predictions on how many genes there are in a certain organism, where they are located, how expression is controlled, and how the genome is organized. It also supports larger questions, like how evolution and specialization occur. Finally, genomic data can be used to study genetic variation within and across species. Tools for analysis of genomic data have been developed and are always being refined. Researchers can now submit several different gene or protein sequences and receive a report that predicts how related the sequences are from an evolutionary point of view. There are also tools to align multiple sequences so we can study how similar and different they are.

DNA fingerprinting

DNA fingerprinting is a laboratory method used to identify individuals based on unique patterns in their genetic code.

-because the DNA of any two people is 99% identical, DNA fingerprinting detects polymorphisms, which are tretches of repetitive and highly variable DNA.

Much like visualizing subtle differences in the whorl patterns of a thumbprint, DNA fingerprinting allows scientists (and police departments!) to detect sequence variations that make each individual’s DNA unique. The ability to appreciate subtle differences within different individual’s DNA comes in handy when matching a DNA sample from a murder suspect to the DNA in a drop of blood found in at a crime scene, or when screening for disease-causing genes, or when doing paternity testing. Since the DNA of any two people is more than 99% identical, DNA fingerprinting exploits stretches of repetitive and highly variable DNA called polymorphisms. These intervening 2-100 base-pair sequences of DNA are structurally variable with respect to their sequence, length, multiplicity, and location within the genome. Two of the several methods of fingerprinting are described below, restriction fragment length polymorphism (RFLP) analysis and short tandem repeat (STR) analysis.

Restriction Fragment Length Polymorphism [RFLP] Analysis

Restriction Fragment Length Polymorphism (RFLP) is a molecular technique that uses restriction enzymes to detect variations (polymorphisms) in homologous DNA sequences.

step 1: This method uses restriction endonucleases to cut 10-100 base pair stretches of polymorphic DNA (called minisatellites) into small fragments. Because of the size variations inherent in this DNA, the resulting DNA fragments (now referred to as RFLPs) also vary in size and are unique to an individual.

Step 2: The RFLPs are separated via gel electrophoresis and transferred to a membrane. Southern blotting techniques are used to analyze the sample. The membrane is probed with radiolabeled DNA oliginucleotides that base-pair specific RFLP sequences, and the mmebrane is visualized with special film. Polymorphic DNA, even though recovered from the same chromosomal region, will yield unique band distributions for each person. When RFLPs are call disease is caused by a single base substition in the beta chain of hemoglobin.

Short Tandem Repeart (STR) Analysis

Short tandem repeat (STR) analysis is a molecular biology method that measures the number of repeating DNA base pairs (STR) at specific locations (loci) in the human genome to create a unique genetic profile.

step 1: this method uses PCR to amplify 5-10 base pair stretches of highly polymorphic and repetitive DNA located within non-coding (introns) regions of the genome. These STRs vary with repsect to the sequence and number of repeats found at each locus. To profile an individual, a smaple of DNA is obtained and the polymorphic DNA is amplified with PCR.

step 2: the amplified STRs are separated via electrophoresis and analyzed with southern blotting (see figure 16)

put figure 16 here.

Additional Methods to Study the Genome

Genomic sequencing is the ultimate study of the genome. However, it is very costly and takes a long time. Depending on the experiment, one of the following methods may be better suited to answering a biological question.

Exome and targeted sequencing

-scientists target certain regions of interest

-exome sequencing: sequencing only the exons of the genome

-on a smaller scale, individual genes can be sequenced

Instead of sequencing the entire genome, scientists can target only certain regions of interest. Exome sequencing involves sequencing only the exons of the genome. On a smaller scale, individual genes can be sequenced. These selective techniques enriching the DNA of interest (by amplification for example), followed by standard sequencing.

Karyotyping

Karyotyping is a laboratory test that examines the size, shape, and number of chromosomes in a cell sample. A normal human karyotype consists of 46 chromosomes organized into 23 pairs: 22 numbered autosomal pairs plus 2 sex chromosomes (XX for female, XY for male).

When generating a karyotype, scientists order all the chromosomes from 1 to 22 plus the sex chromosomes, for a genome-wide view of genetic information. Chromosomes are stained (using Giemsa stain) to highlight structural features. Major genetic changes (involving millions of base pairs), aneuploidy (when a cell contains an abnormal number of chromosomes)

Fluorescence in situ Hybridization

Fluorescence in situ hybridization (FISH) is a molecular cytogenetic technique used to detect and locate specific DNA or RNA sequences on chromosomes or within cells.

Fluorescence in situ hybridization (FISH) uses fluorescently labeled probes to locate the positions of a specific DNA sequences on chromosomes. This detects and localizes the presence or absence of specific DNA sequences on chromosomes. Fluorescence microscopy is used to find out where the fluorescent probe is bound to chromosomes. For example, large chromosomal translocations have been found in several types of cancer. A translocation produces a fusion gene that contains the promoter from one gene and the coding sequence of another. FISH analysis using chromosome 2 and 3 probes can detect and diagnose this translocation.

Analyzing Gene Expression

Many of the techniques discussed above give information about gene expression. For example, RT-PCR and qPCR give information on which genes are being transcribed in a given cell population. Western blot analysis can directly test protein expression, and it is limited only by the amount of starting lysate and the availibility of antibodies specific for the protein being studied. Additional methods have been developed to study gene expression. Each of these techniques can be use to study a certain gene and gather information about its expression and function, or they can be used to to study certain cells and gain information on which genes they are expressing and how they grow and survive.

Microarrays

Microarrays can be used to study relative RNA amounts between two smaples or to compare RNA levels in one sample to normal reference.

This technique is similar to comparitive genomic hybridization (CGH), except that the RNA is used as a starting materail.

1. The two samples of RNA (either two different experimental samples being compared to each other, or one sample and one control) are each labeled with fluorescent dyes of different colors, mixed and applied to an array chip.

2. This chip contains binding sites for every known gene, which act as probes and determine transcript levels in the samples.

3. Figure 17 is comparing gene expression is being compared between two tissue samples (A and B). using a hybridization microarray.

The RNA from sample A is labeled red, and RNA from sample B is labeled green. RNA samples are mixed and applied to teh chip. Genes 1 and 5 are more highly expressed in sample A, since these sites on the chip are red. Genes 2 and 4 are more highly expressed in Sample 5B, since these sites on the chip are green. Gene 3 is expressed in both samples in approximately equal amount resulting in yellow. Microarray data is quantitative, and actual-fold changes can be calculated based on color and intensity

Put figure 17 here

In situ Hybridization

In situ: A Latin phrase meaning "in position" or "in its original place"

hybridization refers to the pairing or annealing of two complementary single-stranded nucleic acid molecules (DNA-DNA, DNA-RNA, or RNA-RNA) to form a double-stranded hybrid structure.

In situ hybridization (ISH) is a laboratory technique used to locate specific DNA or RNA sequences directly within cells, tissue sections, or whole-mount biological samples while preserving tissue architecture.

-ISH is used to determine expression of a gene of interest in a tissue, or in an embryo.

which one is it?

-a very thin slice (or “section”) of a tissue sample is mounted onto a microscope.

-the tissue is fixed to keep transcripts in place

-tissue is permeabilized to open cell membrane

-a labelled probe, specific for the transcript of interest, is added to the section and binds to the transcript being studied

-enzyme-linked antibody is added and binds to the probe

-substrate for enzyme is added

-target-transcript probe antibody complex is detected

in this way, it can be determined when and where transcripts are expressed on a multicellular level

Immunohistochemistry

Immunohistochemistry (IHC) is a laboratory method that uses antibodies to detect specific proteins or antigens in a tissue sample. By combining immunology, histology, and chemistry, IHC allows pathologists to visualize where specific molecules are located within the architecture of a tissue under a microscope.

-similar to ISH, but is specific to proteins instead of nucleic acids

-gives a direct report of protein expression in a tissue

-commonly used in the clinic, for example: breast cancer biopsies from women are stained for the estrogen receptor (ER), the progesterone receptor (PR), and a plasma membrane receptor called (HER2). Breast tumors are then classified as ER⁺ or ER^-, PR⁺, or PR^-, and HER2⁺ or HER2^-. These classifications affect which therapy the patient is given.

1. antibody against a known protein

2. antibody is recognized by a secondary antibody

3. secondary antibody is either linked to an enzyme or fluorescent molecule

look at Figure 18

In figure 18, cells were stained with three different markers.

1. the plasma membrane was labelled green

2. the nucleus was labelled blue

3. the protein of interest was labelled in red

-four different staining patterns were observed

-the two cells at the top express the protein of interest in the cytoplasm

-the three cells in the middle do not express the protein of interest

-the cell at the bottom left expresses the protein of interest in the nucleus

-the blue nuclear stain and red protein of interest are overlapping to make the nucleus purple.

-in the bottom right cell, the protein of interest is expressed on the plasma membrane.

Co-expression of the green plasma membrane marker and the red protein of interest cause yellow staining of the plasma membrane.

Flow Cytometry

Flow cytometry is a laser-based laboratory technique used to measure the physical and chemical characteristics of cells or particles suspended in a fluid.

1. Single cells (either from lab cultures or tissue samples) are stained for certain protein markers using specific antibodies.

2. The antibodies are linked to a fluorescent tag.

3. Next, the labeled cells are suspended in a fluid stream and pass through a beam of light.

4. Light detectors are found on the other side of, and perpendicular to, the laser.

5. As the labelled cells pass through the light, the beam scatters and fluorescent tag(s) on the cells can emit light.

6. This combination of scattered and emitted light is measured to give information on cell size and how many cells in the sample express each of the makers that were labelled. F

Flow cytometeres can have over dozen different light channels, so many labeled antibodies can be added to one experimental sample. In addition to being analyzed, cells can also sorted as they go through the machine (a technique called fluorescence-activated cell sorting, or FACS). In this way, a heterogenous mixture of cells can be sorted based on expression markers.

For example, the earliest (or least differentiated) thymocytes in the thymus express either CD4 nor CD8; therefore, they are classed double negative (CD4^-CD8^-) cells. As these cells progress through their devleopment, they beocme double-positive thermocytes (CD4⁺CD8⁺), and they finally mature to single-positive (CD4⁺CD8^- helper T cells, or CD4^-CD8⁺ killer T cells) thymocytes that are then released from the thymus to peripheral tissues. Each of these four cell populations could be studied using flow cytometry (to determine their relative amounts for example) and could be isolated from each other using FACs.

Protein Quantification

proteonomics: the systematic and large-scale study of protein structure and function.

-involves quantitive analysis of proteins

this means measuring amounts of different proteins from a functional standpoint, looking at how the amount, state, or location of a protein changes. Here are some examples:

-it’s been hypothesized that a particular protein under study functions in G₁ of the cell cycle, but not the other phases.

1. A biochemist tags the protein with a fluorescent molecule, and observes live and cycling cells under fluorescent microscope.

2. He finds that the cells have high levels of fluorescence in G₁, but very low levels of fluorescence in other cell phases.

3. This suggests the protein under study is expressed at high levels in G₁, then is degraded at the beginning of S phase.

-A biochemist is studying the function of an unknown protein, which has been shown to have important functions when a specific transcription is mutated.

1. The biochemist obtains two cell lines.

2. One has a mutation in the transcription factor, and the other doesn’t.

3. She generates lysate samples from the two cell lines [this means she collects and lyses cells, releasing cellular proteins. Other macromolecules (such as lipid bilayers, DNA and RNA) are cleared or degraded by enzymes.

4. She then examines the two lysate sample, looking specifically at the protein of interest.

5. She finds it is not phosphorylated in the cell line without the transcription factor mutation, but is phosphorylated in the cell line with the mutation.

the protein has functions when a specific transcription is mutated, so, therefore, from the results of the study, the protein is fucntioning due to the phosphorylation in the cell line with the mutation.

-a biochemist has a culture of actively growing cells and applies a drug that is being tested for its therapeutic use.

1. the drug targets a protein normally found in the nucleus of the cell.

2. addition of the drug causes the protein to be transported out of the nucleus and into the cytoplasm

3. a quantitative protein experiment measured the total amount of protein in different parts of the cell before and after treatment.

4. It found 90% of the protein in the nucleus before drug treatment, and only 15% was in the nucleus after treatment.

it’s common for proteins to be grown in a biological system, then be extracted and studied. Often, protein levels in lysates or purified samples must be quantified before an experiment can be started. For example, before performing a Western blot, biochemists typically measure protein concentrations in each sample being studied, to make sure the same amount of lysate is loaded into each amount of the gel.

Bradford Quantification

-most used protein quantification method

-uses UV-Vis spectrometers designed for biochemical analysis. This method uses a Bradford reagent containing of a blue pigment called Coomassie blue. When proteins bind the pigment, it shifts the absorption peak of the sample (Figure 19). Absorption is measured at 600nm. This technique is very simple and has good sensitivity.

Put figure 19 here.

to perform bradford protein quantification

1. the negative control is put in the spectrophotometer, to set the zero value

2. samples with known concentration are applied and the spectrophotometer generates a concentration curve

This relates the absorption of the sample with protein concentration

3. a new curve should be made every time proteins are being quantified

4. samples are put in the machine one by one

5. the spectrophotometer applies light in the visible and adjacenet (near-UV and near-infared) ranges.

6. absorbance is determined and compared to the calibration curve, and the machine usualy reports both the absorbance and the subsequent protein concentration.

Affinity chromotography

affinity chromotography is used to separate biochemical mixtures and is based on highly specific interactions betwen macromolecules. While affinty chromotography is most commonly used to purify proteins, it can also be used on other macromolecules (such as nucleic acids). It uses many of the same principeles described above:

1. start with a heterogenous mixture of molecules (such as cell lysate, growth media or bood)

2. to isoalte a protein of interest, you can either use an antibody or tag the protein with an affinity tag (for example, His-tagged proteins can be purified with nickel-based resins and slightly basic conditions; the bound proteins are eluted by adding imidazole or by lowering the pH.)

3. the target molecule is trapped on a stationary phase due to specific binding, and the stationary pahse is washed to increase purity.

4. The target protein isthen released (or eluted) off the solid phase, in a higly purified state.