cells and their sense

Known SNP genotyping.

2008-01-20T10:08:00.000-08:00

This is one of the highly studied methods in the biology field. After the invention of PCR by Mullis a dynamic changes has occurred in this field. Broadly we can classify this in to 2 ,one is conventional PCR RFLP and the other one is multiplexing.

PCR RFLP: In this we will amplify a short segment of DNA which contains our SNP and will cut the amplified DNA with help of an enzyme. These enzymes are known as restriction enzymes. Restriction enzymes are nothing but DNA cutters which cuts DNA at a particular sequence for example EcoR I enzymes the DNA which is having the sequence G/AATTC (/ indicates the site of cut). In this case if we want to see whether GAATTC the A in red is mutated or not. We can need to amplify 100-200 basepairs DNA around this particular sequence and the after amplification we have to incubate the amplicon along with the enzyme. After that we need to run the digested product on to a gel and then we have to see whether the enzyme cut the DNA or not. If the enzyme cuts the DNA means there is no mutation at that particular base and the restriction site is preserved. If there is any mutation the enzyme will not cut the amplified DNA. This method is old fashioned today because it consumes much time when compared to the other applications available. Another restriction is we may not get the enzymes for all the sites which we want to screen.

TaqMan. This application is very rapid and accurate. This method works on the principle that the Taq polymerase have 5’ exonuclease activity. In this we will have a primerset and a probe which is labelled with a Vic and florescent Dye. A Vic blocks the Dyes florescent activity until it is with the probe. If the probe get digested the Vic and Dye will be separated then florescence of the dye will be detected by the system.

In SNP detection we will design a probe in such a condition which will be specific for our SNP and a primer set with the specificity of our (ROI)
this will be further explained in future posts be in touch

Methods to detect SNPs

2008-01-20T09:49:00.000-08:00

There are several methods to SNP identification.
SNP identification can be categorized in to 2 main sub sets.
1. Novel SNP detection.
2. Simply detecting the known SNPs.

For novel snp detection sequencing will be the best method. In this we will select the region which we want to screen for snps from a public database like Ensembl and/or NCBI and we will sequence the region using primers which are specific for that particular region. Once we sequence the region of interest (ROI) we have to compare the sequence to the preexisting sequence to see the variations(SNPs). Here we will get a doubt if we don't have a previous sequence knowledge of that ROI the how can we identify the SNPs. In such case if you are the first person to perform the sequence analysis for that ROI then you have to do the same thing with some other samples after that compare all the sequence to each other and then you will come to know whether there is any variation present in that ROI. To do this we have to use sequence alignment softwares. These softwares align sequences to each others, by doing this we can compare them very fast and very easily. Once we get an idea about the SNp sites in our ROI the we can go for known SNP detection.
Known SNP detection is also known as SNP genotyping. There are several methods available for this. We will discuss about them in future.

If you have any observations please don't hesitate to contact us. Be in touch........

Single Nucleotide Polymorphism(SNP)

2008-01-02T09:42:00.000-08:00

A single nucleotide polymorphism, or SNP (pronounced snip), is a DNA sequence variation occurring when a single nucleotide - A, T, C, or G - in the genome (or other shared sequence) differs between members of a species (or between paired chromosomes in an individual). For example, two sequenced DNA fragments from different individuals, AAGCCTA to AAGCTTA, contain a difference in a single nucleotide. In this case we say that there are two alleles : C and T. Almost all common SNPs have only two alleles.

Within a population, SNPs can be assigned a minor allele frequency - the ratio of chromosomes in the population carrying the less common variant to those with the more common variant. It is important to note that there are variations between human populations, so a SNP allele that is common in one geographical or ethnic group may be much rarer in another. In the past, single nucleotide polymorphisms with a minor allele frequency of greater than or equal to 1% (or 0.5%, etc.) were given the title "SNP," an unwieldy definition. With the advent of modern bioinformatics and a better understanding of evolution, this definition is no longer necessary.

DNA strand 1 differs from DNA strand 2 at a single base-pair location (a C/T polymorphism).

Single nucleotide polymorphisms may fall within coding sequences of genes, non-coding regions of genes, or in the intergenic regions between genes. SNPs within a coding sequence will not necessarily change the amino acid sequence of the protein that is produced, due to degeneracy of the genetic code. A SNP in which both forms lead to the same polypeptide sequence is termed synonymous (sometimes called a silent mutation) - if a different polypeptide sequence is produced they are non-synonymous. SNPs that are not in protein-coding regions may still have consequences for gene splicing, transcription factor binding, or the sequence of non-coding RNA.

Variations in the DNA sequences of humans can affect how humans develop diseases and respond to pathogens, chemicals, drugs, vaccines, and other agents. However, their greatest importance in biomedical research is for comparing regions of the genome between cohorts (such as with matched cohorts with and without a disease).

The study of single nucleotide polymorphisms is also important in crop and livestock breeding programs (see genotyping). See SNP genotyping for details on the various methods used to identify SNPs.

hematopoietic stem cells

2007-10-30T09:25:00.000-07:00

fig: Hematopoietic and Stromal Stem Cell Differentiation.

With more than 50 years of experience studying blood-forming stem cells called hematopoietic stem cells, scientists have developed sufficient understanding to actually use them as a therapy. Currently, no other type of stem cell, adult, fetal or embryonic, has attained such status. Hematopoietic stem cell transplants are now routinely used to treat patients with cancers and other disorders of the blood and immune systems. Recently, researchers have observed in animal studies that hematopoietic stem cells appear to be able to form other kinds of cells, such as muscle, blood vessels, and bone. If this can be applied to human cells, it may eventually be possible to use hematopoietic stem cells to replace a wider array of cells and tissues than once thought.

Scientists face major roadblocks in expanding their use beyond the replacement of blood and immune cells. First, hematopoietic stem cells are unable to proliferate (replicate themselves) and differentiate (become specialized to other cell types) in vitro (in the test tube or culture dish). Second, scientists do not yet have an accurate method to distinguish stem cells from other cells recovered from the blood or bone marrow. Until scientists overcome these technical barriers, they believe it is unlikely that hematopoietic stem cells will be applied as cell replacement therapy in diseases such as diabetes, Parkinson's Disease, spinal cord injury, and many others.

Blood cells are responsible for constant maintenance and immune protection of every cell type of the body. This relentless and brutal work requires that blood cells, along with skin cells, have the greatest powers of self-renewal of any adult tissue.

The stem cells that form blood and immune cells are known as hematopoietic stem cells.They are ultimately responsible for the constant renewal of blood—the production of billions of new blood cells each day. Physicians and basic researchers have known and capitalized on this fact for more than 50 years in treating many diseases. The first evidence and definition of blood-forming stem cells came from studies of people exposed to lethal doses of radiation in 1945.

Basic research soon followed. After duplicating radiation sickness in mice, scientists found they could rescue the mice from death with bone marrow transplants from healthy donor animals. In the early 1960s, Till and McCulloch began analyzing the bone marrow to find out which components were responsible for regenerating blood .They defined what remain the two hallmarks of an HSC: it can renew itself and it can produce cells that give rise to all the different types of blood cells .

A hematopoietic stem cell is a cell isolated from the blood or bone marrow that can renew itself, can differentiate to a variety of specialized cells, can mobilize out of the bone marrow into circulating blood, and can undergo programmed cell death, called apoptosis—a process by which cells that are detrimental or unneeded self-destruct.

A major thrust of basic HSC research since the 1960s has been identifying and characterizing these stem cells. Because HSCs look and behave in culture like ordinary white blood cells, this has been a difficult challenge and this makes them difficult to identify by morphology (size and shape). Even today, scientists must rely on cell surface proteins, which serve, only roughly, as markers of white blood cells.

Identifying and characterizing properties of HSCs began with studies in mice, which laid the groundwork for human studies. The challenge is formidable as about 1 in every 10,000 to 15,000 bone marrow cells is thought to be a stem cell. In the blood stream the proportion falls to 1 in 100,000 blood cells. To this end, scientists began to develop tests for proving the self-renewal and the plasticity of HSCs.

The "gold standard" for proving that a cell derived from mouse bone marrow is indeed an HSC is still based on the same proof described above and used in mice many years ago. That is, the cells are injected into a mouse that has received a dose of irradiation sufficient to kill its own blood-producing cells. If the mouse recovers and all types of blood cells reappear (bearing a genetic marker from the donor animal), the transplanted cells are deemed to have included stem cells.

These studies have revealed that there appear to be two kinds of HSCs. If bone marrow cells from the transplanted mouse can, in turn, be transplanted to another lethally irradiated mouse and restore its hematopoietic system over some months, they are considered to be long-term stem cells that are capable of self-renewal. Other cells from bone marrow can immediately regenerate all the different types of blood cells, but under normal circumstances cannot renew themselves over the long term, and these are referred to as short-term progenitor or precursor cells. Progenitor or precursor cells are relatively immature cells that are precursors to a fully differentiated cell of the same tissue type. They are capable of proliferating, but they have a limited capacity to differentiate into more than one cell type as HSCs do. For example, a blood progenitor cell may only be able to make a red blood cell

Organization and diversity of cells

2007-10-04T13:44:00.000-07:00

All organisms more complex than viruses consist of cells. All cells are derived by cell division from other cells. Ultimately, there must be an unbroken chain of cells leading back to the first successful primordial cell that lived maybe 3.5 billion years ago. How that cell formed is an interesting question.

Prokaryotes lack a defined nucleus and internal organisation is simple. Under the electron microscope they appear featureless. They comprise two kingdoms of life: eubacteria which include most of the bacteria; and the archaea, resemble bacteria and often grow in unusual environments, such as in acid hot springs, saturated brines, etc. The genome of a prokaryote typically consists of a single small circular chromosome in which the DNA is not packaged in any obviously organized way. Prokaryotes may be simple, but they are not primitive .

Fig: Prokaryotic and eukaryotic cell anatomy.

Eukaryotes are thought to have first appeared about 1.5 billion years ago.The organization of the Eukaryotes is complex. Membrane bound- organelles including nucleus are present. Eukaryotic cells have several linear chromosomes in their cell nuclei, in each of which a single very long DNA molecule is elaborately packaged by histone and other proteins. The number and DNA content vary greatly between species .In general the genome size tends to parallel the complexity of the organism, but there are many exceptions. Humans do not have especially large genomes, while the cells of an onion and a lily contain respectively about five and 30 times as much DNA as a typical human cell.

DNA Extraction from Plasma and Serum

2007-09-29T11:03:00.000-07:00

1. Introduction

There are occasions where the only materiel available on a patient is stored plasma or

serum samples. In normal individuals, the amount of DNA in these samples is very low

but sufficient to serve as template for PCRs. Moreover, increased amounts of circulating

DNA have been found in a variety of disorders, including cancer, autoimmune disease,

and infection. Additionally, small amounts of fetal DNA have been detected in maternal

plasma/serum during gestation. We have used the following protocol to successfully

genotype archival plasma samples.

2. Materials

1. 10X SDS /Protein K: (Lauryl sulphate [SDS] 10 g/100 mL, Proteinase K 5 mg/mL).

2. TE (Tris EDTA) buffer: 10 mM Tris-HCl, 1 mM EDTA, pH 8.0.

3. Phenol��chloroform (1��1 v/v).

4. Glycogen (10 mg/mL).

5. 7.5 M Ammonium acetate.

6. 100% ethanol.

7. 70% ethanol.

3. Method

1. Place 1.5 mL of serum or plasma into a 15-mL centrifuge tube.

2. Add 1.5 mL of 1X SDS proteinase K solution in the tube containing the serum and mix

well.

3. Digest overnight at 55°C in water bath.

4. Add 3 mL of phenol/chloroform solution.

5. Vortex 30 s and centrifuge for 10 min at 1000g using a swing-out rotor.

6. Transfer aqueous layer to fresh tube and repeat steps 4 and 5.

7. Transfer aqueous layer to fresh tube and add 5 ìL of glycogen (10 mg/L), 1 mL of 7.5 M

ammonium acetate, and 8 mL of 100% ethanol.

8. Mix by inverting and centrifuge at 2500g for 40 min.

9. Carefully remove supernatant and wash pellet in 10 mL of 70% ethanol.

10. Centrifuge at 2500g for 10 min. Carefully remove last traces of ethanol, and allow to air

dry for 10 min before redissolving in 100 ìL of TE.

RNA Extraction from Blood

2007-09-29T10:45:00.000-07:00

1. Introduction

Based on the method of Chomczynski and Sacchi (1), this is an extremely reliable

method without the requirement for centrifugation over CsCl gradients. As with any

RNA protocol, extreme care should be taken to exclude RNAse contamination, the

greatest source of which will be the sample itself. All disposables and reagents should

be RNAse free.

2. Materials

1. Microfuge tubes (1.5 mL).

2. Ice bucket.

3. Microfuge.

4. Red cell lysis buffer: 1.6 M sucrose, 5% Triton X-100, 25 mM MgCl2, 60 mM Tris-HCl,

pH 7.5; stored at 2–8°C and used cold.

5. Extraction Buffer: 5.25 M guanidinium thiocyanate, 50 mM Tris-Cl, pH. 6.4, 20 mM

EDTA, 1% Triton X-100, 0.1 M â-mercaptoethanol (add immediately prior to use).

6. 2 M sodium acetate, pH 4.0.

7. Phenol (saturated with 1 M Tris-HCl: 0.1 M EDTA, pH 8.0).

8. Chloroform:Iso-amyl alcohol (24��1).

9. Isopropyl alcohol.

10. 70% Ethanol.

11. RNAse-free distilled water.

3. Method

1. In a microfuge tube, mix 100 ìL anticoagulated blood with 1 mL of red cell lysis buffer

(see Notes 1–3).

2. Leave at room temperature with occasional shaking until the red cells have lysed and the

solution translucent (usually within 5 min).

3. Microfuge for 30 s at 13,000g to pellet the white blood cells. Remove and discard

supernatant.

4. Add 200 ìL of extraction buffer and resuspend cell pellet by drawing through narrow

gauge needle several times.

5. Add 20 ìL of 2 M sodium acetate and mix gently by inversion.

6. Add 220 ìL of phenol and mix gently by inversion.

7. Add 60 ìL of chloroform/isoamyl alcohol (24��1) and vortex vigorously.

8. Place on ice for 15 min.

9. Microfuge at 12,000g for 5 min and transfer the upper phase to new microfuge tube.

10. Add 200 ìL of ice-cold isopropanol mix and store at –20°C for 30 min.

11. Microfuge at 12,000g for 15 min and discard supernatant.

12. Resuspend pellet in 200 ìL of extraction buffer.

13. Repeat steps 3 through 9.

14. Wash pellet with 400 ìL of cold 70% ethanol.

15. Microfuge at 12,000g for 5 min and discard supernatant.

16. Carefully remove last traces of ethanol from tube (folded sterile swab or kimwipe works

well).

17. Resuspend in 100 ìL of distilled water and incubate at 50°C for 15 min to dissolve RNA

(see Note 4).

4. Notes

1. Blood stored at room temperature or 4°C should be mixed thoroughly prior to aliquots

being removed.

2. Frozen blood samples should be allowed to thaw completely and mixed thoroughly before

aliquots being removed. Although freezing lyses red blood cells, the red cell lysis step

should still be performed to efficiently remove hemoglobin from the sample. Repeated

freeze/thaw cycles should be avoided.

3. Buffy coat contains two to four times the amount of white blood cells per volume compared

to fresh blood. Therefore, it is advisable to use only 50 ìL of buffy coat diluted with 50 ìL

of phosphate-buffered saline as starting material for this protocol.

4. Repeat pipetting through a narrow gauge tip can help this process.

RT-PCR

2007-09-28T23:28:00.000-07:00

RT-PCR (reverse transcription-polymerase chain reaction) is the most sensitive technique for mRNA detection and quantitation currently available. Compared to the two other commonly used techniques for quantifying mRNA levels, Northern blot analysis and RNase protection assay, RT-PCR can be used to quantify mRNA levels from much smaller samples. In fact, this technique is sensitive enough to enable quantitation of RNA from a single cell.

This article first discusses the advantages of real-time RT-PCR compared to end-point methods. This discussion is followed by a description of the different methods for quantitating gene expression by real-time RT-PCR with respect to the different chemistries available, the quantitation methods used and the instrumentation options available. Subsequently, the “traditional” methods of quantitating gene expression by RT-PCR, i.e. end-point techniques, are presented.

Why Real-Time RT-PCR?

Over the last several years, the development of novel chemistries and instrumentation platforms enabling detection of PCR products on a real-time basis has led to widespread adoption of real-time RT-PCR as the method of choice for quantitating changes in gene expression. Furthermore, real-time RT-PCR has become the preferred method for validating results obtained from array analyses and other techniques that evaluate gene expression changes on a global scale.

To truly appreciate the benefits of real-time PCR, a review of PCR fundamentals is necessary. At the start of a PCR reaction, reagents are in excess, template and product are at low enough concentrations that product renaturation does not compete with primer binding, and amplification proceeds at a constant, exponential rate. The point at which the reaction rate ceases to be exponential and enters a linear phase of amplification is extremely variable, even among replicate samples, but it appears to be primarily due to product renaturation competing with primer binding (since adding more reagents or enzyme has little effect). At some later cycle the amplification rate drops to near zero (plateaus), and little more product is made.

For the sake of accuracy and precision, it is necessary to collect quantitative data at a point in which every sample is in the exponential phase of amplification (since it is only in this phase that amplification is extremely reproducible). Analysis of reactions during exponential phase at a given cycle number should theoretically provide several orders of magnitude of dynamic range. Rare targets will probably be below the limit of detection, while abundant targets will be past the exponential phase. In practice, a dynamic range of 2-3 logs can be quantitated during end-point relative RT-PCR. In order to extend this range, replicate reactions may be performed for a greater or lesser number of cycles, so that all of the samples can be analyzed in the exponential phase.

Real-time PCR automates this otherwise laborious process by quantitating reaction products for each sample in every cycle. The result is an amazingly broad 10⁷-fold dynamic range, with no user intervention or replicates required. Data analysis, including standard curve generation and copy number calculation, is performed automatically. With increasing numbers of labs and core facilities acquiring the instrumentation required for real-time analysis, this technique is becoming the dominant RT-PCR-based quantitation technique.

Real-Time PCR Chemistries

Currently four different chemistries, TaqMan® (Applied Biosystems, Foster City, CA, USA), Molecular Beacons, Scorpions® and SYBR® Green (Molecular Probes), are available for real-time PCR. All of these chemistries allow detection of PCR products via the generation of a fluorescent signal. TaqMan probes, Molecular Beacons and Scorpions depend on Förster Resonance Energy Transfer (FRET) to generate the fluorescence signal via the coupling of a fluorogenic dye molecule and a quencher moeity to the same or different oligonucleotide substrates. SYBR Green is a fluorogenic dye that exhibits little fluorescence when in solution, but emits a strong fluorescent signal upon binding to double-stranded DNA.

TaqMan Probes

TaqMan probes depend on the 5'- nuclease activity of the DNA polymerase used for PCR to hydrolyze an oligonucleotide that is hybridized to the target amplicon. TaqMan probes are oligonucleotides that have a fluorescent reporter dye attached to the 5' end and a quencher moeity coupled to the 3' end. These probes are designed to hybridize to an internal region of a PCR product. In the unhybridized state, the proximity of the fluor and the quench molecules prevents the detection of fluorescent signal from the probe. During PCR, when the polymerase replicates a template on which a TaqMan probe is bound, the 5'- nuclease activity of the polymerase cleaves the probe. This decouples the fluorescent and quenching dyes and FRET no longer occurs. Thus, fluorescence increases in each cycle, proportional to the amount of probe cleavage

Well-designed TaqMan probes require very little optimization. In addition, they can be used for multiplex assays by designing each probe with a spectrally unique fluor/quench pair. However, TaqMan probes can be expensive to synthesize, with a separate probe needed for each mRNA target being analyzed.

Molecular Beacons

Like TaqMan probes, Molecular Beacons also use FRET to detect and quantitate the synthesized PCR product via a fluor coupled to the 5' end and a quench attached to the 3' end of an oligonucleotide substrate. Unlike TaqMan probes, Molecular Beacons are designed to remain intact during the amplification reaction, and must rebind to target in every cycle for signal measurement. Molecular Beacons form a stem-loop structure when free in solution. Thus, the close proximity of the fluor and quench molecules prevents the probe from fluorescing. When a Molecular Beacon hybridizes to a target, the fluorescent dye and quencher are separated, FRET does not occur, and the fluorescent dye emits light upon irradiation.

Molecular Beacons, like TaqMan probes, can be used for multiplex assays by using spectrally separated fluor/quench moieties on each probe. As with TaqMan probes, Molecular Beacons can be expensive to synthesize, with a separate probe required for each target.

Scorpions

With Scorpion probes, sequence-specific priming and PCR product detection is achieved using a single oligonucleotide. The Scorpion probe maintains a stem-loop configuration in the unhybridized state. The fluorophore is attached to the 5' end and is quenched by a moiety coupled to the 3' end. The 3' portion of the stem also contains sequence that is complementary to the extension product of the primer. This sequence is linked to the 5' end of a specific primer via a non-amplifiable monomer. After extension of the Scorpion primer, the specific probe sequence is able to bind to its complement within the extended amplicon thus opening up the hairpin loop. This prevents the fluorescence from being quenched and a signal is observed.

SYBR Green

SYBR Green provides the simplest and most economical format for detecting and quantitating PCR products in real-time reactions. SYBR Green binds double-stranded DNA, and upon excitation emits light. Thus, as a PCR product accumulates, fluorescence increases. The advantages of SYBR Green are that it is inexpensive, easy to use, and sensitive. The disadvantage is that SYBR Green will bind to any double-stranded DNA in the reaction, including primer-dimers and other non-specific reaction products, which results in an overestimation of the target concentration. For single PCR product reactions with well designed primers, SYBR Green can work extremely well, with spurious non-specific background only showing up in very late cycles.

SYBR Green is the most economical choice for real-time PCR product detection. Since the dye binds to double-stranded DNA, there is no need to design a probe for any particular target being analyzed. However, detection by SYBR Green requires extensive optimization. Since the dye cannot distinguish between specific and non-specific product accumulated during PCR, follow up assays are needed to validate results.

Real-time Reporters for Multiplex PCR

TaqMan probes, Molecular Beacons and Scorpions allow multiple DNA species to be measured in the same sample (multiplex PCR), since fluorescent dyes with different emission spectra may be attached to the different probes. Multiplex PCR allows internal controls to be co-amplified and permits allele discrimination in single-tube, homogeneous assays. These hybridization probes afford a level of discrimination impossible to obtain with SYBR Green, since they will only hybridize to true targets in a PCR and not to primer-dimers or other spurious products.

Quantitation of Results

Two strategies are commonly employed to quantify the results obtained by real-time RT-PCR; the standard curve method and the comparative threshold method. These are discussed briefly below.

Standard Curve Method

In this method, a standard curve is first constructed from an RNA of known concentration. This curve is then used as a reference standard for extrapolating quantitative information for mRNA targets of unknown concentrations. Though RNA standards can be used, their stability can be a source of variability in the final analyses. In addition, using RNA standards would involve the construction of cDNA plasmids that have to be in vitro transcribed into the RNA standards and accurately quantitated, a time-consuming process. However, the use of absolutely quantitated RNA standards will help generate absolute copy number data.

In addition to RNA, other nucleic acid samples can be used to construct the standard curve, including purified plasmid dsDNA, in vitro generated ssDNA or any cDNA sample expressing the target gene. Spectrophotometric measurements at 260 nm can be used to assess the concentration of these DNAs, which can then be converted to a copy number value based on the molecular weight of the sample used. cDNA plasmids are the preferred standards for standard curve quantitation. However, since cDNA plasmids will not control for variations in the efficiency of the reverse transcription step, this method will only yield information on relative changes in mRNA expression. This, and variation introduced due to variable RNA inputs, can be corrected by normalization to a housekeeping gene.

Comparative C_t Method

Another quantitation approach is termed the comparative C_t method. This involves comparing the C_t values of the samples of interest with a control or calibrator such as a non-treated sample or RNA from normal tissue. The C_t values of both the calibrator and the samples of interest are normalized to an appropriate endogenous housekeeping gene.

The comparative C_t method is also known as the 2^{–[delta][delta]Ct} method, where

[delta][delta]C_t = [delta]C_t,sample - [delta]C_t,reference

Here, [delta]C_T,sample is the C_t value for any sample normalized to the endogenous housekeeping gene and [delta]C_{t, reference} is the C_t value for the calibrator also normalized to the endogenous housekeeping gene.

For the [delta][delta]C_t calculation to be valid, the amplification efficiencies of the target and the endogenous reference must be approximately equal. This can be established by looking at how [delta]C_t varies with template dilution. If the plot of cDNA dilution versus delta C_t is close to zero, it implies that the efficiences of the target and housekeeping genes are very similar. If a housekeeping gene cannot be found whose amplification efficiency is similar to the target, then the standard curve method is preferred.

Instrumentation for Real-Time PCR

Real-time PCR requires an instrumentation platform that consists of a thermal cycler, a computer, optics for fluorescence excitation and emission collection, and data acquisition and analysis software. These machines, available from several manufacturers, differ in sample capacity (some are 96-well standard format, others process fewer samples or require specialized glass capillary tubes), method of excitation (some use lasers, others broad spectrum light sources with tunable filters), and overall sensitivity. There are also platform-specific differences in how the software processes data. Real-time PCR machines are not inexpensive, currently about $25K - $95K, but are well within purchasing reach of core facilities or labs that have the need for high throughput quantitative analysis. For a comprehensive list of real-time thermal cyclers please see the weblink at the end of this article.

Tools for Real-Time RT-PCR

Ambion’s MessageSensor™ RT Kit includes an RNase H+ MMLV RT that clearly outperforms MMLV RT enzymes that have abolished RNase H activity in real-time RT-PCR experiments. Unlike many other qRT-PCR kits, MessageSensor includes a total RNA control, a control human GAPDH primer set, RNase inhibitor, and nucleotides, as well as a buffer additive that enables detection with SYBR® Green dye.

The Cells-to-cDNA™ II Kit produces cDNA from cultured mammalian cells in less than 2 hours. No RNA isolation is required. This kit is ideal for those who want to perform reverse transcription reactions on small numbers of cells, numerous cell samples, or for scientists who are unfamiliar with RNA isolation. Ambion's Cells-to-cDNA II Kit contains a novel Cell Lysis Buffer that inactivates endogenous RNases without compromising downstream enzymatic reactions. After inactivation of RNases, the cell lysate can be directly added to a cDNA synthesis reaction. Cells-to-cDNA II is compatible with both one-step and two-step real-time RT-PCR protocols.

Genomic DNA contamination can lead to false positive RT-PCR results. Ambion offers a variety of tools for eliminating genomic DNA contamination from RNA samples prior to RT-PCR. Ambion’s DNA-free™ DNase Treatment and Removal Reagents are designed for removing contaminating DNA from RNA samples and for the removal of DNase after treatment without Proteinase K treatment and organic extraction. In addition, Ambion has also developed TURBO™ DNase, a hyperactive enzyme engineered from wild-type bovine DNase. The proficiency of TURBO DNase in binding very low concentrations of DNA means that the enzyme is particularly effective in removing trace quantities of DNA contamination.

Ambion now also offers an economical alternative to the high cost of PCR reagents for the ABI 7700 and other 0.2 ml tube-based real-time instruments. SuperTaq™ Real-Time performs as well or better than the more expensive alternatives, and includes dNTPs and a Reaction Buffer optimized for SYBR Green, TaqMan, and Molecular Beacon chemistries.

End-Point RT-PCR: Relative vs. Competitive vs. Comparative

In spite of the rapid advances made in the area of real-time PCR detection chemistries and instrumentation, end-point RT-PCR still remains a very commonly used technique for measuring changes in gene-expression in small sample numbers.

End-point RT-PCR can be used to measure changes in expression levels using three different methods: relative, competitive and comparative. The most commonly used procedures for quantitating end-point RT-PCR results rely on detecting a fluorescent dye such as ethidium bromide, or quantitation of P³²-labeled PCR product by a phosphorimager or, to a lesser extent, by scintillation counting.

Relative quantitation compares transcript abundance across multiple samples, using a co-amplified internal control for sample normalization. Results are expressed as ratios of the gene-specific signal to the internal control signal. This yields a corrected relative value for the gene-specific product in each sample. These values may be compared between samples for an estimate of the relative expression of target RNA in the samples; for example, 2.5-fold more IL-12 in sample 2 than in sample 1.

Absolute quantitation, using competitive RT-PCR, measures the absolute amount (e.g., 5.3 x 10⁵ copies) of a specific mRNA sequence in a sample. Dilutions of a synthetic RNA (identical in sequence, but slightly shorter than the endogenous target) are added to sample RNA replicates and are co-amplified with the endogenous target. The PCR product from the endogenous transcript is then compared to the concentration curve created by the synthetic "competitor RNA."

Comparative RT-PCR mimics competitive RT-PCR in that target message from each RNA sample competes for amplification reagents within a single reaction, making the technique reliably quantitative. Because the cDNA from both samples have the same PCR primer binding site, one sample acts as a competitor for the other, making it unnecessary to synthesize a competitor RNA sequence.

Both relative and competitive RT-PCR quantitation techniques require pilot experiments. In the case of relative RT-PCR, pilot experiments include selection of a quantitation method and determination of the exponential range of amplification for each mRNA under study. For competitive RT-PCR, a synthetic RNA competitor transcript must be synthesized and used in pilot experiments to determine the appropriate range for the standard curve. Comparative RT-PCR yields similar sensitivity as relative and competitive RT-PCR, but requires significantly less optimization and does not require synthesis of a competitor.

Relative RT-PCR

Relative RT-PCR uses primers for an internal control that are multiplexed in the same RT-PCR reaction with the gene specific primers. Internal control and gene-specific primers must be compatible — that is, they must not produce additional bands or hybridize to each other. The expression of the internal control should be constant across all samples being analyzed. Then the signal from the internal control can used to normalize sample data to account for tube-to-tube differences caused by variable RNA quality or RT efficiency, inaccurate quantitation or pipetting. Common internal controls include ß-actin and GAPDH mRNAs and 18S rRNA. Unlike Northerns and nuclease protection assays, where an internal control probe is simply added to the experiment, the use of internal controls in relative RT-PCR requires substantial optimization.

For relative RT-PCR data to be meaningful, the PCR reaction must be terminated when the products from both the internal control and the gene of interest are detectable and are being amplified within exponential phase (see Determining Exponential Range in PCR). Because internal control RNAs are typically constituitively expressed housekeeping genes of high abundance, their amplification surpasses exponential phase with very few PCR cycles. It is therefore difficult to identify compatible exponential phase conditions where the PCR product from a rare message is detectable. Detection methods with low sensitivity, like ethidium bromide staining of agarose gels, are therefore not recommended. Detecting a rare message while staying in exponential range with an abundant message can be achieved several ways: 1) by increasing the sensitivity of product detection, 2) by decreasing the amount of input template in the RT or PCR reactions and/or 3) by decreasing the number of PCR cycles.

Ambion recommends using 18S rRNA as an internal control because it shows less variance in expression across treatment conditions than ß-actin and GAPDH. However, because of its abundance, it is difficult to detect the PCR product for rare messages in the exponential phase of amplification of 18S rRNA. Ambion's patented Competimer™ Technology solves this problem by attenuating the 18S rRNA signal even to the level of rare messages. Attenuation results from the use of competimers — primers identical in sequence to the functional 18S rRNA primers but that are "blocked" at their 3'-end and, thus, cannot be extended by PCR. Competimers and primers are mixed at various ratios to reduce the amount of PCR product generated from 18S rRNA. Figure 1 illustrates that 18S rRNA primers without competimers cannot be used as an internal control because the 18S rRNA amplification overwhelms that of clathrin (compare panels A and B). Mixing primers with competimers at a 3:7 ratio attenuates the 18S rRNA signal, making 18S rRNA a practical internal control (panel C).

Figure 1. Ambion's QuantumRNA™ Technology in Multiplex Quantitative RT-PCR using 18S rRNA as an Internal Control. RT-PCR reactions on brain, embryo, liver, and spleen total RNA using A) primers for clathrin, B) primers for clathrin and 18S, or C) primers for clathrin, 18S rRNA primers and 18S rRNA Competimers. Note that without Competimers, 18S cannot be used as an internal control because of its high abundance (B). Addition of Competimers (C) makes multiplex PCR possible, providing sample-to-sample relative quantitation.

Ambion's QuantumRNA 18S Internal Standards contain 18S rRNA primers and competimers designed to amplify 18S rRNA in all eukaryotes. The Universal 18S Internal Standards function across the broadest range of organisms including plants, animals and many protozoa. The Classic I and Classic II 18S Internal Standards can be used with any vertebrate RNA sample. All 18S Internal Standards work well in multiplex RT-PCR. These kits also include control RNA and an Instruction Manual detailing the series of experiments needed to make relative RT-PCR data significant. For those researchers who have validated ß-actin as an appropriate internal control for their system, the QuantumRNA ß-actin Internal Standards are available.

Competitive RT-PCR

Competitive RT-PCR precisely quantitates a message by comparing RT-PCR product signal intensity to a concentration curve generated by a synthetic competitor RNA sequence. The competitor RNA transcript is designed for amplification by the same primers and with the same efficiency as the endogenous target. The competitor produces a different-sized product so that it can be distinguished from the endogenous target product by gel analysis. The competitor is carefully quantitated and titrated into replicate RNA samples. Pilot experiments are used to find the range of competitor concentration where the experimental signal is most similar. Finally, the mass of product in the experimental samples is compared to the curve to determine the amount of a specific RNA present in the sample.

Some protocols use DNA competitors or random sequences for competitive RT-PCR. These competitors do not effectively control for variations in the RT reaction or for the amplification efficiency of the specific experimental sequence, as do RNA competitors. See The Accuracy of Competitive RT-PCR Depends on Using the Right Exogenous Standard for a further discussion on competitor choice and design.

Comparative RT-PCR

While exquisitely sensitive, both relative and competitive methods of qRT-PCR have drawbacks. Relative RT-PCR requires extensive optimization to ensure that the PCR is terminated when both the gene of interest and an internal control are in the exponential phase of amplification. Competitive RT-PCR requires that an exogenous "competitor" be synthesized for each target to be analyzed. However, comparative RT-PCR achieves the same level of sensitivity as these standard methods of qRT-PCR, with significantly less optimization. Target mRNAs from 2 samples are assayed simultaneously, each serving as a competitor for the other, making it possible to compare the relative abundance of target between samples. Comparative RT-PCR is ideal for analyzing target genes discovered by screening methods such as array analysis and differential display.

Tools for Any RT-PCR Technique

Whether you choose to perform real-time, relative, competitive, or comparative RT-PCR, Ambion offers products to simplify your RT-PCR experiments and make the data more quantitative. In addition to the specific products described above, Ambion offers SuperTaq™ Polymerase, M-MLV Reverse Transcriptase, and RNase-free PCR tubes. To prevent cross contamination during PCR experiments, Ambion also offers DNAZap™ DNA Degradation Solution and RNase-free barrier pipette tips.

For a comprehensive list of publications discussing practically every aspect of real-time RT-PCR please visit www.wzw.tum.de/gene-quantification/real-time.html