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Instrumentation and Techniques -Aliasgar Vohra

Sr.no Topic Page.no 1. Flow Cytometry 1-3 2. Fluorescence Activated Cell-Sorting 4-7 3. Cell-cycle Analysis 8-9 4. Polymerase Chain Reaction(PCR) 10-14 5. Real-time Polymerase Chain Reaction(RT-PCR) 15-22 6. Polymerase Chain Reaction Stages 23-24 7. Types of PCR 25-30 8. Variants of PCR 31-35 9. Primer Modification 35-36

10. DNA Polymerase 37-39 11. GEL Electrophoresis 40-47 12. Macrophage 48-55 13. Electrophoresis 56-57 14. Cell-Culture 58-64 15. Common Cell Lines 65 16. Types of Cell Cultures 66 17. Basic Cell Cultures Techniques 67 18. Growth Medium 68-72 19. Basic Constituents of Media 73-77 20. Laminar Flow 78-79 21. Laminar Flow Reactor 80-81 22. Incubator 82 23. Fluorescence Microscope 83-86 24. ELISA 87-89 25. Types of ELISA 90-96 26. Four Typical ELISA Format 97-100

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Flow cytometry In biotechnology, flow cytometry is a laser-based, biophysical technology employed in cell counting, cell sorting, biomarker detection and protein engineering, by suspendingcells in a stream of fluid and passing them by an electronic detection apparatus. It allows simultaneous multiparametric analysis of the physical and chemical characteristics of up to thousands of particles per second.

Flow cytometry is routinely used in the diagnosis of health disorders, especially blood cancers, but has many other applications in basic research, clinical practice and clinical trials. A common variation is to physically sort particles based on their properties, so as to purify populations of interest.

History The first impedance-based flow cytometry device, using the Coulter principle, was disclosed in U.S. Patent 2,656,508, issued in 1953, to Wallace H. Coulter. Mack Fulwyler was the inventor of the forerunner to today's flow cytometers - particularly the cell sorter.[1 ] Fulwyler developed this in 1965 with his publication in Science.[2 ] The first fluorescence-based flow cytometry device (ICP 11) was developed in 1968 by Wolfgang Göhde from the University of Münster, filed for patent on 18 December 1968[3] and first commercialized in 1968/69 by German developer and manufacturer Partec through Phywe AG in Göttingen. At that time, absorption methods were still widely favored by other scientists over fluorescence methods.[4 ] Soon after, flow cytometry instruments were developed, including the Cytofluorograph (1971) from Bio/Physics Systems Inc. (later: Ortho Diagnostics), the PAS 8000 (1973) from Partec, the first FACS (Fluorescence-activated cell sorting) instrument from Becton Dickinson (1974), the ICP 22 (1975) from Partec/Phywe and the Epics from Coulter (1977/78).

Name of the technology The original name of the fluorescence-based flow cytometry technology was "pulse cytophotometry" (German: Impulszytophotometrie), based on the first patent application on fluorescence-based flow cytometry. At the 5th American Engineering Foundation Conference on Automated Cytology in Pensacola (Florida) in 1976 - eight years after the introduction of the first fluorescence-based flow cytometer (1968) - it was agreed to commonly use the name "flow cytometry", a term that quickly became popular.[5]

Flow cytometers

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Front view of a desktop flow cytometer - the Becton-DickinsonFluorescence activated cell sorter (FACSCalibur)

Modern flow cytometers are able to analyze several thousand particles every second, in "real time," and can actively separate and isolate particles having specified properties. A flow cytometer is similar to a microscope, except that, instead of producing an image of the cell, flow cytometry offers "high-throughput" (for a large number of cells) automated quantification of set parameters. To analyze solid tissues, a single-cell suspension must first be prepared.

A flow cytometer has five main components:

a flow cell - liquid stream (sheath fluid), which carries and aligns the cells so that they pass

single file through the light beam for sensing

a measuring system - commonly used are measurement of impedance (or conductivity) and

optical systems - lamps (mercury, xenon); high-power water-cooled lasers (argon, krypton, dye

laser); low-power air-cooled lasers (argon (488 nm), red-HeNe (633 nm), green-HeNe, HeCd

(UV)); diode lasers (blue, green, red, violet) resulting in light signals

a detector and Analogue-to-Digital Conversion (ADC) system - which generates forward-

scattered light (FSC) and side-scattered light (SSC) as well as fluorescence signals from light

into electrical signals that can be processed by a computer

an amplification system - linear or logarithmic

a computer for analysis of the signals.

The process of collecting data from samples using the flow cytometer is termed 'acquisition'. Acquisition is mediated by a computer physically connected to the flow cytometer, and the software which handles the digital interface with the cytometer. The software is capable of adjusting parameters (e.g., voltage, compensation) for the sample being tested, and also assists in displaying initial sample information while acquiring sample data to ensure that parameters are set correctly. Early flow cytometers were, in general, experimental devices, but technological advances have enabled widespread applications for use in a variety of both clinical and research purposes. Due to these developments, a considerable market for instrumentation, analysis software, as well as the reagents used in acquisition such as fluorescently labeled antibodies has developed.

Modern instruments usually have multiple lasers and fluorescence detectors. The current record for a commercial instrument is ten lasers[6] and 18 fluorescence detectors. Increasing the number of lasers and detectors allows for multiple antibody labeling, and can more precisely identify a target population by their phenotypic markers. Certain instruments can even take digital images of individual cells, allowing for the analysis of fluorescent signal location within or on the surface of cells.

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Data analysis

Analysis of a marine sample of photosyntheticpicoplankton by flow cytometry showing three different populations

(Prochlorococcus, Synechococcus, andpicoeukaryotes)

Gating The data generated by flow-cytometers can be plotted in a single dimension, to produce a histogram, or in two-dimensional dot plots or even in three dimensions. The regions on these plots can be sequentially separated, based on fluorescenceintensity, by creating a series of subset extractions, termed "gates." Specific gating protocols exist for diagnostic and clinical purposes especially in relation to hematology.

The plots are often made on logarithmic scales. Because different fluorescent dyes' emission spectra overlap,[7][8 ] signals at the detectors have to be compensated electronically as well as computationally. Data accumulated using the flow cytometer can be analyzed using software, e.g., WinMDI,[9] Flowing Software,[10] and web-based Cytobank[11] (all freeware), FCS Express, Flowjo, FACSDiva, CytoPaint (aka Paint-A-Gate),[12] VenturiOne, CellQuest Pro, Infinicyt or Cytospec.[13] Once the data are collected, there is no need to stay connected to the flow cytometer. For this reason, analysis is most often performed on a separate computer. This is especially necessary in core facilities where usage of these machines is in high demand.

Computational analysis Recent progress on automated population identification using computational methods has offered an alternative to traditional gating strategies. Automated identification systems could potentially help findings of rare and hidden populations. Representative automated methods include FLOCK [14] in Immunology Database and Analysis Portal (ImmPort),[15] SamSPECTRAL[16] and flowClust[17][18][1 9] in Bioconductor, and FLAME [20] in GenePattern. Collaborative efforts have resulted in an open project called FlowCAP (Flow Cytometry: Critical Assessment of Population Identification Methods,[21]) to provide an objective way to compare and evaluate the flow cytometry data clustering methods, and also to establish guidance about appropriate use and application of these methods.

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Fluorescence-activated cell sorting (FACS)

Fluorescence-Activated Cell Sorting (FACS)

Fluorescence-activated cell sorting (FACS) is a specialized type of flow cytometry. It provides a method for sorting a heterogeneous mixture of biological cells into two or more containers, one cell at a time, based upon the specific light scattering and fluorescent characteristics of each cell. It is a useful scientific instrument as it provides fast, objective and quantitative recording of fluorescent signals from individual cells as well as physical separation of cells of particular interest. The acronym FACS is trademarked and owned by Becton, Dickinson and Company.[22 ] Among the large majority of researchers who use this technology for sorting or analysis, this term has become generic in common usage, much like xerox or kleenex. The first cell sorter was invented by Mack Fulwyler in 1965, using the Coulter principle, a relatively difficult technique that is no longer used in modern instruments. The technique was expanded by Len Herzenberg, who was responsible for coining the term FACS.[2 3] Herzenberg won the Kyoto Prize in 2006 for his seminal work in flow cytometry.

The cell suspension is entrained in the center of a narrow, rapidly flowing stream of liquid. The flow is arranged so that there is a large separation between cells relative to their diameter. A vibrating mechanism causes the stream of cells to break into individual droplets. The system is adjusted so that there is a low probability of more than one cell per droplet. Just before the stream breaks into droplets, the flow passes through a fluorescence measuring station where the fluorescent character of interest of each cell is measured. An electrical charging ring is placed just at the point where the stream breaks into droplets. A charge is placed on the ring based on the immediately prior fluorescence intensity measurement, and the opposite charge is trapped on the droplet as it breaks from the stream. The charged droplets then fall through anelectrostatic deflection system that diverts droplets into containers based upon their charge. In some systems, the charge is applied directly to the stream, and the droplet breaking off retains charge of the same sign as the stream. The stream is then returned to neutral after the droplet breaks off.

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Labels

Use of flow cytometry to measure copy number variation of a specific DNA sequence (Flow-FISH)

Fluorescent labels A wide range of fluorophores can be used as labels in flow cytometry.[24] Fluorophores, or simply "fluors", are typically attached to an antibody that recognises a target feature on or in the cell; they may also be attached to a chemical entity with affinity for the cell membrane or another cellular structure. Each fluorophore has a characteristic peak excitation and emissionwavelength, and the emission spectra often overlap. Consequently, the combination of labels which can be used depends on the wavelength of the lamp(s) or laser(s) used to excite the fluorochromes and on the detectors available.[25] The maximum number of distinguishable fluorescent labels is thought to be 17 or 18, and this level of complexity necessitates laborious optimization to limit artifacts, as well as complex deconvolution algorithms to separate overlapping spectra.[26 ] Besides flow cytometry is a quantitative mean of fluorescence, the utmost sensitivity of flow cytometry is unmatched by other fluorescent detection platforms such as confocal microscopy. Absolute fluorescence sensitivity is generally lower in confocal microscopybecause out-of-focus signals are rejected by the confocal optical system and because the image is built up serially from individual measurements at every location across the cell, reducing the amount of time available to collect signal.[27]

Quantum dots Quantum dots are sometimes used in place of traditional fluorophores because of their narrower emission peaks.

Isotope labeling Main article: Mass cytometry

Mass cytometry overcomes the fluorescent labeling limit by utilizing lanthanide isotopes attached to antibodies. This method could theoretically allow the use of 40 to 60 distinguishable labels and has been demonstrated for 30 labels.[26 ] Mass cytometry is fundamentally different from flow cytometry: cells are introduced into a plasma, ionized, and associated isotopes are quantified via time-of-flight mass spectrometry. Although this method permits the use of a large number of labels, it currently has lower throughput capacity than flow cytometry. It also destroys the analysed cells, precluding their recovery by sorting.[26]

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Cytometric Bead Array (CBA) In addition to the ability to label and identify individual cells via fluorescent antibodies, cellular products such as cytokines, proteins, and other factors may also be measured as well. Similar to ELISA sandwich assays, CBA assays use multiple bead populations typically differentiated by size and different levels of fluorescence intensity to distinguish multiple analytes in a single assay. The amount of the analyte captured is detected via a biotinylated antibody against a secondary epitope of the protein, followed by a streptavidin-R-phycoerythrin treatment. The fluorescent intensity of R-phycoerythrin on the beads is quantified on a flow cytometer equipped with a 488 nm excitation source. Concentrations of a protein of interest in the samples can be obtained by comparing the fluorescent signals to those of a standard curve generated from a serial dilution of a known concentration of the analyte. Commonly also referred to as cytokine bead array (CBA).

Measurable parameters This list is very long and constantly expanding.

used for confirming diagnosis of chronic lymphocytic leukemia

volume and morphological complexity of cells

cell pigments such as chlorophyll or phycoerythrin

total DNA content (cell cycle analysis,

cell kinetics, proliferation, ploidy, aneuploidy, endoreduplication, etc.)

total RNA content

DNA copy number variation (by Flow-FISH or BACs-on-Beads technology)

chromosome analysis and sorting (library construction, chromosome paint)

protein expression and localization

protein modifications, phospho-proteins

transgenic products in vivo, particularly the Green fluorescent protein or related

Fluorescent Proteins

cell surface antigens (Cluster of differentiation (CD) markers)

intracellular antigens (various cytokines, secondary mediators, etc.)

nuclear antigens

enzymatic activity

pH, intracellular ionized calcium, magnesium, membrane potential

membrane fluidity

apoptosis (quantification, measurement of DNA degradation, mitochondrial membrane

potential, permeability changes, caspase activity)

cell viability

monitoring electropermeabilization of cells

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oxidative burst

characterising multidrug resistance (MDR) in cancer cells

glutathione

various combinations (DNA/surface antigens, etc.)

cell adherence (for instance pathogen-host cell adherence)

Applications The technology has applications in a number of fields, including molecular biology, pathology, immunology, plant biology and marine biology.[28] It has broad application inmedicine (especially in transplantation, hematology, tumor immunology and chemotherapy, prenatal diagnosis, genetics and sperm sorting for sex preselection). Also, it is extensively used in research for the detection of DNA damage, caspase cleavage and apoptosis.[29] In marine biology, the autofluorescent properties of photosynthetic planktoncan be exploited by flow cytometry in order to characterise abundance and community structure. In protein engineering, flow cytometry is used in conjunction with yeast displayand bacterial display to identify cell surface-displayed protein variants with desired properties.

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Cell-cycle analysis Cell-cycle analysis is a method in cell biology that employs flow cytometry to distinguish cells in different phases of the cell cycle. Before analysis, the cells are permeabilisedand treated with a fluorescent dye that stains DNA quantitatively, usually propidium iodide (PI). The fluorescence intensity of the stained cells at certain wavelengths will therefore correlate with the amount of DNA they contain. As the DNA content of cells duplicates during the S phase of the cell cycle, the relative amount of cells in the G0 phase and G1phase (before S phase), in the S phase, and in the G2 phase and M phase (after S phase) can be determined, as the fluorescence of cells in the G2/M phase will be twice as high as that of cells in the G0/G1 phase.

Cell-cycle anomalies can be symptoms for various kinds of cell damage, for example DNA damage, which cause the cell to interrupt the cell cycle at certain checkpoints to prevent transformation into a cancer cell (carcinogenesis). Other possible reasons for anomalies include lack of nutrients, for example after serum deprivation.

Cell cycle analysis was first described in 1969 at Los Alamos Scientific Laboratory by a group from the University of California,[1] using the Feulgen staining technique. The first protocol for cell-cycle analysis using propidium iodide staining was presented in 1975 by Awtar Krishan from Harvard Medical School[2] and is still widely cited today.

Experimental procedure

DAPI (magenta) bound to the minor groove of DNA (green and blue). From PDB1D30.

The first step in preparing cells for cell-cycle analysis is permeabilisation of the cells' plasma membranes. This is usually done by incubating them in a buffer solution containing a detergent [3] such as Triton X-100 or NP-40, or by fixating them in ethanol. Most fluorescent DNA dyes are not membrane permeable, that is, unable to pass through an intact cell membrane. Permeabilisation is therefore crucial for the success of the next step, the staining of the cells.

Prior to or during the staining step, the cells will usually be treated with RNase A to remove RNAs from the cells. This is important because dyes that stain DNA will also stain RNA, thus creating artefacts that would distort the results.

Aside from propidium iodide, quantifiable dyes that are frequently used include (but are not limited to) DRAQ5, 7-Aminoactinomycin D, DAPI and Hoechst 33342.

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When the cells pass through the flow cytometer's laser, a fluorescence pulse is generated that correlates with the amount of dye associated with the DNA and thus with the total amount of DNA in the cell.

Doublet discrimination Because cells and especially fixated cells tend to stick together, cell aggregates have to be excluded from analysis through a process called doublet discrimination. This is important because a doublet of two cells in the G0/G1 phase has the same total amount of DNA and thus the same fluorescence intensity as a single cell in the G2/M phase.[4 ]G0/G1 doublets would therefore create false positive results for G2/M cells.

Related Methods Nicoletti assay

The Nicoletti assay, named after its inventor, the Italian physician Ildo Nicoletti, is a modified form of cell cycle analysis. It is used to detect and measure apoptosis, a form ofprogrammed cell death, by analysing cells with a DNA content less than 2n ("sub-G1 cells"). Such cells are usually the result of apoptotic DNA fragmentation: during apoptosis, the DNA is degraded by cellular endonucleases. Therefore, nuclei of apoptotic cells contain less DNA than nuclei of healthy G0/G1 cells, resulting in a sub-G0/G1 peak in the fluorescence histogram that can be used to determine the relative amount of apoptotic cells in a sample.

This method was developed and first described in 1991 by Nicoletti and co-workers at Perugia University School of Medicine.[5] An optimised protocol developed by two of the authors of the original publication was published in 2006. [6]

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Polymerase chain reaction

A strip of eight PCR tubes, each containing a 100 μl reaction mixture

The polymerase chain reaction (PCR) is a technology in molecular biology used to amplify a single copy or a few copies of a piece of DNA across several orders of magnitude, generating thousands to millions of copies of a particular DNA sequence.

Developed in 1983 by Kary Mullis,[1][2 ] PCR is now a common and often indispensable technique used in medical and biological research labs for a variety of applications.[3][4 ] These include DNA cloning for sequencing, DNA-based phylogeny, or functional analysis of genes; the diagnosis of hereditary diseases; the identification of genetic fingerprints (used in forensic sciences and paternity testing); and the detection and diagnosis of infectious diseases. In 1993, Mullis was awarded theNobel Prize in Chemistry along with Michael Smith for his work on PCR.[5]

The method relies on thermal cycling, consisting of cycles of repeated heating and cooling of the reaction for DNA meltingand enzymatic replication of the DNA. Primers (short DNA fragments) containing sequences complementary to the target region along with a DNA polymerase, which the method is named after, are key components to enable selective and repeated amplification. As PCR progresses, the DNA generated is itself used as a template for replication, setting in motion a chain reaction in which the DNA template is exponentially amplified. PCR can be extensively modified to perform a wide array ofgenetic manipulations.

Almost all PCR applications employ a heat-stable DNA polymerase, such as Taq polymerase (an enzyme originally isolated from the bacterium Thermus aquaticus). This DNA polymerase enzymatically assembles a new DNA strand from DNA building-blocks, the nucleotides, by using single-stranded DNA as a template and DNA oligonucleotides (also called DNA primers), which are required for initiation of DNA synthesis. The vast majority of PCR methods use thermal cycling, i.e., alternately heating and cooling the PCR sample through a defined series of temperature steps.

In the first step, the two strands of the DNA double helix are physically separated at a high temperature in a process called DNA melting. In the second step, the temperature is lowered and the two DNA strands become templates for DNA polymerase to selectively amplify the target DNA.

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The selectivity of PCR results from the use of primers that arecomplementary to the DNA region targeted for amplification under specific thermal cycling conditions.

Placing a strip of eight PCR tubes, each containing a 100 μl reaction mixture, into the thermal cycler

PCR principles and procedure

Figure 1a: A thermal cycler for PCR

Figure 1b: An older model three-temperature thermal cycler for PCR

PCR amplifies a specific region of a DNA strand (the DNA target). Most PCR methods typically amplify DNA fragments of between 0.1 and 10kilo base pairs (kbp), although some techniques allow for amplification of fragments up to 40 kbp in size. [6] The amount of amplified product is

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determined by the available substrates in the reaction, which become limiting as the reaction progresses.[7]

A basic PCR set up requires several components and reagents.[8 ] These components include:

DNA template that contains the DNA region (target) to amplify

Two primers that are complementary to the 3' (three prime) ends of each of the sense and

anti-sense strand of the DNA target

Taq polymerase or another DNA polymerase with a temperature optimum at around 70 °C

Deoxynucleoside triphosphates (dNTPs, sometimes called "deoxynucleotide

triphosphates"; nucleotides containing triphosphate groups), the building-blocks from

which the DNA polymerase synthesizes a new DNA strand

Buffer solution, providing a suitable chemical environment for optimum activity and

stability of the DNA polymerase

Bivalent cations, magnesium or manganese ions; generally Mg2+ is used, but Mn2+ can be

used for PCR-mediated DNA mutagenesis, as higher Mn2 + concentration increases the error

rate during DNA synthesis[9]

Monovalent cation potassium ions

The PCR is commonly carried out in a reaction volume of 10–200 μl in small reaction tubes (0.2–0.5 ml volumes) in a thermal cycler. The thermal cycler heats and cools the reaction tubes to achieve the temperatures required at each step of the reaction (see below). Many modern thermal cyclers make use of the Peltier effect, which permits both heating and cooling of the block holding the PCR tubes simply by reversing the electric current. Thin-walled reaction tubes permit favorable thermal conductivity to allow for rapid thermal equilibration. Most thermal cyclers have heated lids to prevent condensation at the top of the reaction tube. Older thermocyclers lacking a heated lid require a layer of oil on top of the reaction mixture or a ball of wax inside the tube.

Procedure Typically, PCR consists of a series of 20-40 repeated temperature changes, called cycles, with each cycle commonly consisting of 2-3 discrete temperature steps, usually three (Figure below). The cycling is often preceded by a single temperature step at a high temperature (>90 °C), and followed by one hold at the end for final product extension or brief storage. The temperatures used and the length of time they are applied in each cycle depend on a variety of parameters. These include the enzyme used for DNA synthesis, the concentration of divalent ions and dNTPs in the reaction, and the melting temperature (Tm) of the primers. [10]

Initialization step(Only required for DNA polymerases that require heat activation

by hot-start PCR.[11]): This step consists of heating the reaction to a temperature of 94–96 °C (or

98 °C if extremely thermostable polymerases are used), which is held for 1–9 minutes.

Denaturation step: This step is the first regular cycling event and consists of heating

the reaction to 94–98 °C for 20–30 seconds. It causes DNA melting of the DNA template by

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disrupting the hydrogen bonds between complementary bases, yielding single-stranded DNA

molecules.

Annealing step: The reaction temperature is lowered to 50–65 °C for 20–40 seconds

allowing annealing of the primers to the single-stranded DNA template. This temperature must

be low enough to allow for hybridization of the primer to the strand, but high enough for the

hybridization to be specific, i.e., the primer should only bind to a perfectly complementary part

of the template. If the temperature is too low, the primer could bind imperfectly. If it is too high,

the primer might not bind. Typically the annealing temperature is about 3–5 °C below the Tm of

the primers used. Stable DNA–DNA hydrogen bonds are only formed when the primer sequence

very closely matches the template sequence. The polymerase binds to the primer-template

hybrid and begins DNA formation.

Extension/elongation step: The temperature at this step depends on the DNA

polymerase used; Taq polymerase has its optimum activity temperature at 75–80 °C,[12 ][13] and

commonly a temperature of 72 °C is used with this enzyme. At this step the DNA polymerase

synthesizes a new DNA strand complementary to the DNA template strand by adding dNTPs

that are complementary to the template in 5' to 3' direction, condensing the 5'-phosphate

group of the dNTPs with the 3'-hydroxyl group at the end of the nascent (extending) DNA

strand. The extension time depends both on the DNA polymerase used and on the length of the

DNA fragment to amplify. As a rule-of-thumb, at its optimum temperature, the DNA polymerase

polymerizes a thousand bases per minute. Under optimum conditions, i.e., if there are no

limitations due to limiting substrates or reagents, at each extension step, the amount of DNA

target is doubled, leading to exponential (geometric) amplification of the specific DNA

fragment.

Final elongation: This single step is occasionally performed at a temperature of 70–

74 °C (this is the temperature needed for optimal activity for most polymerases used in PCR)

for 5–15 minutes after the last PCR cycle to ensure that any remaining single-stranded DNA is

fully extended.

Final hold: This step at 4–15 °C for an indefinite time may be employed for short-term

storage of the reaction.

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Figure 3: Ethidium bromide-stained PCR products after gel electrophoresis. Two sets of primers were used to

amplify a target sequence from three different tissue samples. No amplification is present in sample #1; DNA bands

in sample #2 and #3 indicate successful amplification of the target sequence. The gel also shows a positive control,

and a DNA ladder containing DNA fragments of defined length for sizing the bands in the experimental PCRs.

To check whether the PCR generated the anticipated DNA fragment (also sometimes referred to as the amplimer oramplicon), agarose gel electrophoresis is employed for size separation of the PCR products. The size(s) of PCR products is determined by comparison with a DNA ladder (a molecular weight marker), which contains DNA fragments of known size, run on the gel alongside the PCR products (see Fig. 3).

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Real-time polymerase chain reaction From Wikipedia, the free encyclopedia For reverse transcription polymerase chain reaction (RT-PCR), see reverse transcription polymerase chain reaction.

SYBR Green fluorescence chart for five samples, each having three replicates, which is a result of quantitative PCR

(qPCR).

Melting curve for five samples, three replicates each, which is a result of melting temperature analysis of

quantitative PCR results (qPCR).

A real-time polymerase chain reaction is a laboratory technique of molecular biology based on the polymerase chain reaction (PCR), which is used to amplify and simultaneously detect or quantify a targeted DNA molecule.

The procedure follows the general principle of polymerase chain reaction; its key feature is that the amplified DNA is detected as the reaction progresses in "real time". This is a new approach compared to standard PCR, where the product of the reaction is detected at its end. Two common methods for the detection of products in quantitative PCR are: (1) non-specificfluorescent dyes that intercalate with any double-stranded DNA, and (2) sequence-specific DNA probes consisting ofoligonucleotides that are labelled with a fluorescent reporter which permits detection only after hybridization of the probe with its complementary sequence to quantify messenger RNA (mRNA) and non-coding RNA in cells or tissues.

The MIQE guidelines propose that the abbreviation qPCR be used for quantitative real-time PCR and that RT-qPCR be used for reverse transcription–qPCR [1]. The acronym "RT-PCR" commonly denotes reverse transcription polymerase chain reaction and not real-time PCR, but not all authors adhere to this convention.[1]

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Background

Real time quantitative PCR uses fluorophores in order to detect levels of gene expression.

Cells in all organisms regulate gene expression by turnover of gene transcripts (messenger RNA, abbreviated to mRNA): The amount of an expressed gene in a cell can be measured by the number of copies of an mRNA transcript of that gene present in a sample. In order to robustly detect and quantify gene expression from small amounts of RNA, amplification of the gene transcript is necessary. The polymerase chain reaction (PCR) is a common method for amplifying DNA; for mRNA-based PCR the RNA sample is first reverse-transcribed to cDNA with reverse transcriptase.

In order to amplify small amounts of DNA, the same methodology is used as in conventional PCR using a DNA template, at least one pair of specific primers, deoxyribonucleotides, a suitable buffer solution and a thermo-stable DNA polymerase. A substance marked with a fluorophore is added to this mixture in a thermal cycler that contains sensors for measuring thefluorescence of the flurophore after it has been excited at the required wavelength allowing the generation rate to be measured for one or more specific products. This allows the rate of generation of the amplified product to be measured at each PCR cycle. The data thus generated can be analysed by computer software to calculate relative gene expression (ormRNA copy number) in several samples. Quantitative PCR can also be applied to the detection and quantification of DNA in samples to determine the presence and abundance of a particular DNA sequence in these samples.[2] This measurement is made after each amplification cycle, and this is the reason why this method is called real time PCR (that is, immediate or simultaneous PCR). In the case of RNA quantitation, the template is complementary DNA (cDNA), which is obtained by reverse transcription of ribonucleic acid (RNA). In this instance the technique used is quantitative RT-PCR or Q-RT-PCR.

Quantitative PCR and DNA microarray are modern methodologies for studying gene expression. Older methods were used to measure mRNA abundance: Differential display,RNase protection assay and Northern blot. Northern blotting is often used to estimate the expression level of a gene by visualizing the abundance of its mRNA transcript in a sample. In this method, purified RNA is separated by agarose gel electrophoresis, transferred to a solid matrix (such as a nylon membrane), and probed with a specific DNA or RNA probe that is complementary to the gene of interest. Although this technique is still used to assess gene expression, it requires relatively large amounts of RNA and provides only qualitative or semi quantitative information of mRNA levels.[3] Estimation errors arising from variations in the quantification method can be the result of DNA integrity, enzyme efficiency and many other factors. For this reason a number of standardization systems have been developed. Some have been developed for quantifying total gene expression, but the most common are aimed at quantifying the specific gene being studied in

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relation to another gene called a normalizing gene, which is selected for its almost constant level of expression. These genes are often selected from housekeeping genes as their functions related to basic cellular survival normally implie constitutive gene expression.[4 ][5] This enables researchers to report a ratio for the expression of the genes of interest divided by the expression of the selected normalizer, thereby allowing comparison of the former without actually knowing its absolute level of expression.

The most commonly used normalizing genes are those that code for the following molecules: tubulin, glyceraldehyde-3-phosphate dehydrogenase, albumin, cyclophilin, andribosomal RNAs.[3]

Basic principles Quantitative PCR is carried out in a thermal cycler with the capacity to illuminate each sample with a beam of light of a specified wavelength and detect the fluorescence emitted by the excited fluorophore. The thermal cycler is also able to rapidly heat and chill samples, thereby taking advantage of the physicochemical properties of the nucleic acids andDNA polymerase.

The PCR process generally consists of a series of temperature changes that are repeated 25 – 40 times. These cycles normally consist of three stages: the first, at around 95 °C, allows the separation of the nucleic acid’s double chain; the second, at a temperature of around 50-60 °C, allows the binding of the primers with the DNA template; [6] the third, at between 68 - 72 °C, facilitates the polymerization carried out by the DNA polymerase. Due to the small size of the fragments the last step is usually omitted in this type of PCR as the enzyme is able to increase their number during the change between the alignment stage and the denaturing stage. In addition, some thermal cyclers add another short temperature phase lasting only a few seconds to each cycle, with a temperature of, for example, 80 °C, in order to reduce the noise caused by the presence of primer dimers when a non-specific dye is used. The temperatures and the timings used for each cycle depend on a wide variety of parameters, such as: the enzyme used to synthesize the DNA, the concentration of divalent ions and deoxyribonucleotides (dNTPs) in the reaction and the bonding temperature of the primers. [7]

Classification The type of quantitative PCR technique used depends on the DNA sequence in the samples, the technique can either use non-specific fluorochromes or hybridization probes.

Quantitative PCR with double-stranded DNA-binding dyes as reporters

A DNA-binding dye binds to all double-stranded (ds) DNA in PCR, causing fluorescence of the dye. An increase in DNA product during PCR therefore leads to an increase in fluorescence intensity and is measured at each cycle, thus allowing DNA concentrations to be quantified. However, dsDNA dyes such as SYBR Green will bind to all dsDNA PCR products, including nonspecific PCR products (such as Primer dimer). This can potentially interfere with, or prevent, accurate quantification of the intended target sequence. The SYBR Green is excited using blue light (λmax = 488 nm) and it emits green light (λmax = 522 nm).[8]

The reaction is prepared as usual, with the addition of fluorescent dsDNA dye.

The reaction is run in a quantitative PCR instrument, and after each cycle, the levels of

fluorescence are measured with a detector; the dye only fluoresces when bound to the

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dsDNA (i.e., the PCR product). With reference to a standard dilution, the dsDNA

concentration in the PCR can be determined.

This method has the advantage of only needing a pair of primers to carry out the amplification, which keeps costs down; however, it is only possible to amplify a product using a chain reaction.

Like other quantitative PCR methods, the values obtained do not have absolute units associated with them (i.e., mRNA copies/cell). As described above, a comparison of a measured DNA/RNA sample to a standard dilution will only give a fraction or ratio of the sample relative to the standard, allowing only relative comparisons between different tissues or experimental conditions. To ensure accuracy in the quantification, it is usually necessary to normalize expression of a target gene to a stably expressed gene (see below). This can correct possible differences in RNA quantity or quality across experimental samples.

Fluorescent reporter probe method

(1) In intact probes, reporter fluorescence is quenched. (2) Probes and the complementary DNA strand are

hybridized and reporter fluorescence is still quenched. (3) During PCR, the probe is degraded by the Taq

polymerase and the fluorescent reporter released.

Fluorescent reporter probes detect only the DNA containing the probe sequence; therefore, use of the reporter probe significantly increases specificity, and enables quantification even in the presence of non-specific DNA amplification. Fluorescent probes can be used in multiplex assays—for detection of several genes in the same reaction—based on specific probes with different-coloured labels, provided that all targeted genes are amplified with similar efficiency. The specificity of fluorescent reporter probes also prevents interference of measurements caused byprimer dimers, which are undesirable potential by-products in PCR. However, fluorescent reporter probes do not prevent the inhibitory effect of the primer dimers, which may depress accumulation of the desired products in the reaction.

The method relies on a DNA-based probe with a fluorescent reporter at one end and a quencher of fluorescence at the opposite end of the probe. The close proximity of the reporter to the quencher prevents detection of its fluorescence; breakdown of the probe by the 5' to 3' exonuclease activity of the Taq polymerase breaks the reporter-quencher proximity and thus allows unquenched emission of fluorescence, which can be detected after excitation with a laser. An increase in the product targeted by the reporter probe at each PCR cycle therefore causes a proportional increase in fluorescence due to the breakdown of the probe and release of the reporter.

The PCR is prepared as usual (see PCR), and the reporter probe is added.

20

As the reaction commences, during the annealing stage of the PCR both probe and

primers anneal to the DNA target.

Polymerisation of a new DNA strand is initiated from the primers, and once the

polymerase reaches the probe, its 5'-3'-exonuclease degrades the probe, physically

separating the fluorescent reporter from the quencher, resulting in an increase in

fluorescence.

Fluorescence is detected and measured in a real-time PCR machine, and its geometric

increase corresponding to exponential increase of the product is used to determine the

quantification cycle (Cq) in each reaction.

Fusion temperature analysis

Distinct fusion curves for a number of PCR products (showing distinct colours). Amplification reactions can be seen

for a specific product (pink, blue) and others with a negative result (green, orange). The fusion peak indicated with

an arrow shows the peak caused by primer dimers, which is different from the expected amplification product.[9]

Q-PCR permits the identification of specific, amplified DNA fragments using analysis of their melting temperature (also called Tm value, from melting temperature). The method used is usually PCR with double-stranded DNA-binding dyes as reporters and the dye used is usually SYBR Green. The DNA melting temperature is specific to the amplified fragment. The results of this technique are obtained by comparing the dissociation curves of the analysed DNA samples.[10 ]

Unlike conventional PCR, this method avoids the previous use of electrophoresis techniques to demonstrate the results of all the samples. This is because, despite being a kinetic technique, quantitative PCR is usually evaluated at a distinct end point. The technique therefore usually provides more rapid results and / or uses fewer reactants than electrophoresis. If subsequent electrophoresis is required it is only necessary to test those samples that real time PCR has shown to be doubtful and / or to ratify the results for samples that have tested positive for a specific determinant.

21

Quantification of gene expression

Quantifying gene expression by traditional DNA detection methods is unreliable. Detection of mRNA on a Northern blot or PCR products on a gel or Southern blot does not allow precise quantification.[11] For example, over the 20-40 cycles of a typical PCR, the amount of DNA product reaches a plateau that is not directly correlated with the amount of target DNA in the initial PCR.[citation needed]

Quantitative PCR can be used to quantify nucleic acids by two common methods: relative quantification and absolute quantification.[12]Absolute quantification gives the exact number of target DNA molecules by comparison with DNA standards using a calibration curve. It is therefore essential that the PCR of the sample and the standard have the same amplification efficiency. Relative quantification is based on internal reference genes to determine fold-differences in expression of the target gene. The quantification is expressed as the change in expression levels of mRNA interpreted as complementary DNA (cDNA, generated by reverse transcription of mRNA). Relative quantification is easier to carry out as it does not require a calibration curve as the amount of the studied gene is compared to the amount of a control housekeeping gene.

As the units used to express the results of relative quantification are unimportant the results can be compared across a number of different RT-Q-PCR. The reason for using one or more housekeeping genes is to correct non-specific variation, such as the differences in the quantity and quality of RNA used, which can affect the efficiency of reverse transcription and therefore that of the whole PCR process. However, the most crucial aspect of the process is that the reference gene must be stable.[13]

The selection of these reference genes was traditionally carried out in molecular biology using qualitative or semi-quantitative studies such as the visual examination of RNA gels, Northern blot densitometry or semi-quantitative PCR (PCR mimics). Now, in the genome era, it is possible to carry out a more detailed estimate for many organisms using DNA microarrays.[1 4] However, research has shown that amplification of the majority of reference genes used in quantifying the expression of mRNA varies according to experimental conditions.[15][16][17] It is therefore necessary to carry out an initial statistically sound methodological study in order to select the most suitable reference gene.

A number of statistical algorithms have been developed that can detect which gene or genes are most suitable for use under given conditions. Those like geNORM or BestKeeper can compare pairs or geometric means for a matrix of different reference genes and tissues.[18][1 9]

Modeling

Unlike end point PCR (conventional PCR) real time PCR allows quantification of the desired product at any point in the amplification process by measuring fluorescence (in reality, measurement is made of its level over a given threshold). A commonly employed method of DNA quantification by quantitative PCR relies on plotting fluorescence against the number of cycles on a logarithmic scale. A threshold for detection of DNA-based fluorescence is set slightly above background. The number of cycles at which the fluorescence exceeds the threshold is called the threshold cycle (Ct) or, according to the MIQE guidelines, quantification cycle (Cq).[20]

During the exponential amplification phase, the quantity of the target DNA template (amplicon) doubles every cycle. For example, a DNA sample whose Cq precedes that of another sample by 3 cycles contained 23 = 8 times more template. However, the efficiency of amplification is often variable among primers and templates. Therefore, the efficiency of a primer-template combination is assessed in a titration experiment with serial dilutions of DNA template to create a standard

22

curve of the change in Cq with each dilution. The slope of the linear regression is then used to determine the efficiency of amplification, which is 100% if a dilution of 1:2 results in a Cq difference of 1. The cycle threshold method makes several assumptions of reaction mechanism and has a reliance on data from low signal-to-noise regions of the amplification profile that can introduce substantial variance during the data analysis.[21]

To quantify gene expression, the Cq for an RNA or DNA from the gene of interest is subtracted from the Cq of RNA/DNA from a housekeeping gene in the same sample to normalize for variation in the amount and quality of RNA between different samples. This normalization procedure is commonly called the ΔCt-method[22] and permits comparison of expression of a gene of interest among different samples. However, for such comparison, expression of the normalizing reference gene needs to be very similar across all the samples. Choosing a reference gene fulfilling this criterion is therefore of high importance, and often challenging, because only very few genes show equal levels of expression across a range of different conditions or tissues.[23 ][24] Although cycle threshold analysis is integrated with many commercial software systems, there are more accurate and reliable methods of analysing amplification profile data that should be considered in cases where reproducibility is a concern.[21]

Mechanism-based qPCR quantification methods have also been suggested, and have the advantage that they do not require a standard curve for quantification. Methods such as MAK2[25] have been shown to have equal or better quantitative performance to standard curve methods. These mechanism-based methods use knowledge about the polymerase amplification process to generate estimates of the original sample concentration. An extension of this approach includes an accurate model of the entire PCR reaction profile, which allows for the use of high signal-to-noise data and the ability to validate data quality prior to analysis.[21]

Applications There are numerous applications for quantitative polymerase chain reaction in the laboratory. It is commonly used for both diagnostic and basic research. Uses of the technique in industry include the quantification of microbial load in foods or on vegetable matter, the detection of GMOs (Genetically modified organisms) and the quantification and genotyping of human viral pathogens.

Diagnostic uses

Diagnostic quantitative PCR is applied to rapidly detect nucleic acids that are diagnostic of, for example, infectious diseases, cancer and genetic abnormalities. The introduction of quantitative PCR assays to the clinical microbiology laboratory has significantly improved the diagnosis of infectious diseases,[2 6] and is deployed as a tool to detect newly emerging diseases, such as new strains of flu, in diagnostic tests.[27]

Microbiological uses

Quantitative PCR is also used by microbiologists working in the fields of food safety, food spoilage and fermentation and for the microbial risk assessment of water quality (drinking and recreational waters) and in public health protection.[28]

The antibacterial assay Virtual Colony Count[29 ] utilizes a data quantification technique called Quantitative Growth Kinetics (QGK) that is mathematically identical to QPCR, except bacterial cells, rather than copies of a PCR product, increase exponentially. The QGK equivalent of the threshold cycle is referred to as the "threshold time".

23

Uses in research

In research settings, quantitative PCR is mainly used to provide quantitative measurements of gene transcription. The technology may be used in determining how the genetic expression of a particular gene changes over time, such as in the response of tissue and cell cultures to an administration of a pharmacological agent, progression of cell differentiation, or in response to changes in environmental conditions. It is also used for the determination of zygosity of transgenic animals used in research.

Detection of phytopathogens

The agricultural industry is constantly striving to produce plant propagules or seedlings that are free of pathogens in order to prevent economic losses and safeguard health. Systems have been developed that allow detection of small amounts of the DNA of Phytophthora ramorum, an oomycete that kills Oaks and other species, mixed in with the DNA of the host plant. Discrimination between the DNA of the pathogen and the plant is based on the amplification of ITS sequences, spacers located in ribosomal RNA gene’s coding area, which are characteristic for each taxon.[30] Field-based versions of this technique have also been developed for identifying the same pathogen.[31 ]

Detection of genetically modified organisms

qPCR using reverse transcription (RT-qPCR) can be used to detect GMOs given its sensitivity and dynamic range in detecting DNA. Alternatives such as DNA or protein analysis are usually less sensitive. Specific primers are used that amplify not the transgene but the promoter, terminator or even intermediate sequences used during the process of engineering the vector. As the process of creating a transgenic plant normally leads to the insertion of more than one copy of the transgene its quantity is also commonly assessed. This is often carried out by relative quantification using a control gene from the treated species that is only present as a single copy.

Clinical quantification and genotyping

Viruses can be present in humans due to direct infection or co-infections. This makes diagnosis difficult using classical techniques and can result in an incorrect prognosis and treatment. The use of qPCR allows both the quantification and genotyping (characterization of the strain, carried out using melting curves) of a virus such as the Hepatitis B virus.[34 ] The degree of infection, quantified as the copies of the viral genome per unit of the patient’s tissue, is relevant in many cases; for example, the probability that the type 1herpes simplex virus reactivates is related to the number of infected neurons in the ganglia.[35 ] This quantification is carried out either with reverse transcription or without it, as occurs if the virus becomes integrated in the human genome at any point in its cycle, such as happens in the case of HPV (human papillomavirus), where some of its variants are associated with the appearance of cervical cancer.[36]

24

PCR stages

The PCR process can be divided into three stages:

Exponential amplification: At every cycle, the amount of product is doubled (assuming 100% reaction efficiency). The reaction is very sensitive: only minute quantities of DNA must be present.[1 4]

Leveling off stage: The reaction slows as the DNA polymerase loses activity and as consumption of reagents such as dNTPs and primers causes them to become limiting.

Plateau: No more product accumulates due to exhaustion of reagents and enzyme.

PCR optimization In practice, PCR can fail for various reasons, in part due to its sensitivity to contamination causing amplification of spurious DNA products. Because of this, a number of techniques and procedures have been developed for optimizing PCR conditions. [15][16] Contamination with extraneous DNA is addressed with lab protocols and procedures that separate pre-PCR mixtures from potential DNA contaminants.[8] This usually involves spatial separation of PCR-setup areas from areas for analysis or purification of PCR products, use of disposable plasticware, and thoroughly cleaning the work surface between reaction setups. Primer-design techniques are important in improving PCR product yield and in avoiding the formation of spurious products, and the usage of alternate buffer components or polymerase enzymes can help with amplification of long or otherwise problematic regions of DNA. Addition of reagents, such as formamide, in buffer systems may increase the specificity and yield of PCR.[17] Computer simulations of theoretical PCR results (Electronic PCR) may be performed to assist in primer design.[18 ]

Application of PCR Selective DNA isolation

PCR allows isolation of DNA fragments from genomic DNA by selective amplification of a specific region of DNA. This use of PCR augments many methods, such as generatinghybridization probes for Southern or northern hybridization and DNA cloning, which require larger amounts of DNA, representing a specific DNA region. PCR supplies these techniques with high amounts of pure DNA, enabling analysis of DNA samples even from very small amounts of starting material.

Other applications of PCR include DNA sequencing to determine unknown PCR-amplified sequences in which one of the amplification primers may be used in Sanger sequencing, isolation of a DNA sequence to expedite recombinant DNA technologies involving the insertion of a DNA sequence into a plasmid, phage, or cosmid (depending on size) or the genetic material of another organism. Bacterial colonies (E. coli) can be rapidly screened by PCR for correct DNA vector constructs.[19] PCR may also be used forgenetic fingerprinting; a forensic technique used to identify a person or organism by comparing experimental DNAs through different PCR-based methods.

Some PCR 'fingerprints' methods have high discriminative power and can be used to identify genetic relationships between individuals, such as parent-child or between siblings, and are used in paternity testing (Fig. 4). This technique may also be used to determine evolutionary relationships among organisms when certain molecular clocks are used (i.e., the 16S rRNA and recA genes of microorganisms).[citation needed]

25

Figure 4: Electrophoresis of PCR-amplified DNA fragments. (1) Father. (2) Child. (3) Mother. The child has

inherited some, but not all of the fingerprint of each of its parents, giving it a new, unique fingerprint.

Amplification and quantification of DNA Because PCR amplifies the regions of DNA that it targets, PCR can be used to analyze extremely small amounts of sample. This is often critical for forensic analysis, when only a trace amount of DNA is available as evidence. PCR may also be used in the analysis of ADNA that is tens of thousands of years old. These PCR-based techniques have been successfully used on animals, such as a forty-thousand-year-oldmammoth, and also on human DNA, in applications ranging from the analysis of Egyptian mummies to the identification of a Russian tsar and the body of English king Richard III.[20]

Quantitative PCR methods allow the estimation of the amount of a given sequence present in a sample—a technique often applied to quantitatively determine levels of gene expression. Quantitative PCR is an established tool for DNA quantification that measures the accumulation of DNA product after each round of PCR amplification.

PCR in diagnosis of diseases PCR permits early diagnosis of malignant diseases such as leukemia and lymphomas, which is currently the highest-developed in cancer research and is already being used routinely. PCR assays can be performed directly on genomic DNA samples to detect translocation-specific malignant cells at a sensitivity that is at least 10,000 fold higher than that of other methods.[c itation needed]

PCR allows for rapid and highly specific diagnosis of infectious diseases, including those caused by bacteria or viruses.[21 ] PCR also permits identification of non-cultivatable or slow-growing microorganisms such as mycobacteria, anaerobic bacteria, or viruses from tissue cultureassays and animal models. The basis for PCR diagnostic applications in microbiology is the detection of infectious agents and the discrimination of non-pathogenic from pathogenic strains by virtue of specific genes.[21][22]

26

Viral DNA can likewise be detected by PCR. The primers used must be specific to the targeted sequences in the DNA of a virus, and the PCR can be used for diagnostic analyses or DNA sequencing of the viral genome. The high sensitivity of PCR permits virus detection soon after infection and even before the onset of disease.[21] Such early detection may give physicians a significant lead time in treatment. The amount of virus ("viral load") in a patient can also be quantified by PCR-based DNA quantitation techniques (see below).

Limitations

DNA polymerase is prone to error, which in turn causes mutations in the PCR fragments that are made. Additionally, the specificity of the PCR fragments can mutate to the template DNA, due to nonspecific binding of primers. Furthermore prior information on the sequence is necessary in order to generate the primers. [23]

Variations on the basic PCR technique

Allele-specific PCR: a diagnostic or cloning technique based on single-nucleotide

variations (SNVs not to be confused with SNPs) (single-base differences in a patient). It

requires prior knowledge of a DNA sequence, including differences between alleles, and uses

primers whose 3' ends encompass the SNV (base pair buffer around SNV usually incorporated).

PCR amplification under stringent conditions is much less efficient in the presence of a

mismatch between template and primer, so successful amplification with an SNP-specific

primer signals presence of the specific SNP in a sequence.[2 4] See SNP genotyping for more

information.

Assembly PCR or Polymerase Cycling Assembly (PCA): artificial synthesis of

long DNA sequences by performing PCR on a pool of long oligonucleotides with short

overlapping segments. The oligonucleotides alternate between sense and antisense directions,

and the overlapping segments determine the order of the PCR fragments, thereby selectively

producing the final long DNA product.[25]

Asymmetric PCR: preferentially amplifies one DNA strand in a double-stranded DNA

template. It is used in sequencing and hybridization probing where amplification of only one of

the two complementary strands is required. PCR is carried out as usual, but with a great excess

of the primer for the strand targeted for amplification. Because of the slow (arithmetic)

amplification later in the reaction after the limiting primer has been used up, extra cycles of

PCR are required. [26] A recent modification on this process, known as Linear-After-The-

Exponential-PCR (LATE-PCR), uses a limiting primer with a higher melting temperature (Tm)

than the excess primer to maintain reaction efficiency as the limiting primer concentration

decreases mid-reaction.[27]

Dial-out PCR: a highly parallel method for retrieving accurate DNA molecules for gene

synthesis. A complex library of DNA molecules is modified with unique flanking tags before

27

massively parallel sequencing. Tag-directed primers then enable the retrieval of molecules with

desired sequences by PCR.[28 ]

Digital PCR (dPCR): used to measure the quantity of a target DNA sequence in a DNA

sample. The DNA sample is highly diluted so that after running many PCRs in parallel, some of

them do not receive a single molecule of the target DNA. The target DNA concentration is

calculated using the proportion of negative outcomes. Hence the name 'digital PCR'.

Helicase-dependent amplification: similar to traditional PCR, but uses a constant

temperature rather than cycling through denaturation and annealing/extension cycles. DNA

helicase, an enzyme that unwinds DNA, is used in place of thermal denaturation.[2 9]

Hot start PCR: a technique that reduces non-specific amplification during the initial set up

stages of the PCR. It may be performed manually by heating the reaction components to the

denaturation temperature (e.g., 95 °C) before adding the polymerase.[30 ] Specialized enzyme

systems have been developed that inhibit the polymerase's activity at ambient temperature,

either by the binding of an antibody[11][31 ] or by the presence of covalently bound inhibitors that

dissociate only after a high-temperature activation step. Hot-start/cold-finish PCR is achieved

with new hybrid polymerases that are inactive at ambient temperature and are instantly

activated at elongation temperature.

In silico PCR (digital PCR, virtual PCR, electronic PCR, e-PCR) refers to

computational tools used to calculate theoretical polymerase chain reaction results using a

given set of primers (probes) to amplify DNA sequences from a

sequenced genome or transcriptome. In silico PCR was proposed as an educational tool for

molecular biology.[32]

Intersequence-specific PCR (ISSR): a PCR method for DNA fingerprinting that

amplifies regions between simple sequence repeats to produce a unique fingerprint of

amplified fragment lengths. [33]

Inverse PCR: is commonly used to identify the flanking sequences around genomic inserts.

It involves a series of DNA digestions and self ligation, resulting in known sequences at either

end of the unknown sequence.[3 4]

Ligation-mediated PCR: uses small DNA linkers ligated to the DNA of interest and

multiple primers annealing to the DNA linkers; it has been used for DNA sequencing,genome

walking, and DNA footprinting.[35 ]

Methylation-specific PCR (MSP): developed by Stephen Baylin and Jim Herman at the

Johns Hopkins School of Medicine,[36] and is used to detect methylation of CpG islands in

genomic DNA. DNA is first treated with sodium bisulfite, which converts unmethylated cytosine

bases to uracil, which is recognized by PCR primers as thymine. Two PCRs are then carried out

28

on the modified DNA, using primer sets identical except at any CpG islands within the primer

sequences. At these points, one primer set recognizes DNA with cytosines to amplify

methylated DNA, and one set recognizes DNA with uracil or thymine to amplify unmethylated

DNA. MSP using qPCR can also be performed to obtain quantitative rather than qualitative

information about methylation.

Miniprimer PCR: uses a thermostable polymerase (S-Tbr) that can extend from short

primers ("smalligos") as short as 9 or 10 nucleotides. This method permits PCR targeting to

smaller primer binding regions, and is used to amplify conserved DNA sequences, such as the

16S (or eukaryotic 18S) rRNA gene. [37]

Multiplex Ligation-dependent Probe Amplification (MLPA): permits

amplifying multiple targets with a single primer pair, thus avoiding the resolution limitations of

multiplex PCR (see below).

Multiplex-PCR: consists of multiple primer sets within a single PCR mixture to

produce amplicons of varying sizes that are specific to different DNA sequences. By targeting

multiple genes at once, additional information may be gained from a single test-run that

otherwise would require several times the reagents and more time to perform. Annealing

temperatures for each of the primer sets must be optimized to work correctly within a single

reaction, and amplicon sizes. That is, their base pair length should be different enough to form

distinct bands when visualized by gel electrophoresis.

Nanoparticle-Assisted PCR (nanoPCR): In recent years, it has been reported that

some nanoparticles (NPs) can enhance the efficiency of PCR (thus being called nanoPCR), and

some even perform better than the original PCR enhancers. It was also found that quantum dots

(QDs) can improve PCR specificity and efficiency. Single-walled carbon nanotubes (SWCNTs)

and multi-walled carbon nanotubes (MWCNTs) are efficient in enhancing the amplification of

long PCR. Carbon nanopowder (CNP) was reported be able to improve the efficiency of

repeated PCR and long PCR. ZnO, TiO2, and Ag NPs were also found to increase PCR yield.

Importantly, already known data has indicated that non-metallic NPs retained acceptable

amplification fidelity. Given that many NPs are capable of enhancing PCR efficiency, it is clear

that there is likely to be great potential for nanoPCR technology improvements and product

development.[38][39]

Nested PCR: increases the specificity of DNA amplification, by reducing background due to

non-specific amplification of DNA. Two sets of primers are used in two successive PCRs. In the

first reaction, one pair of primers is used to generate DNA products, which besides the intended

target, may still consist of non-specifically amplified DNA fragments. The product(s) are then

used in a second PCR with a set of primers whose binding sites are completely or partially

29

different from and located 3' of each of the primers used in the first reaction. Nested PCR is

often more successful in specifically amplifying long DNA fragments than conventional PCR, but

it requires more detailed knowledge of the target sequences.

Overlap-extension PCR or Splicing by overlap extension (SOEing) :

a genetic engineering technique that is used to splice together two or more DNA fragments that

contain complementary sequences. It is used to join DNA pieces containing genes, regulatory

sequences, or mutations; the technique enables creation of specific and long DNA constructs. It

can also introduce deletions, insertions or point mutations into a DNA sequence.[40 ][41 ]

PAN-AC: uses isothermal conditions for amplification, and may be used in living cells.[42 ][43]

quantitative PCR (qPCR): used to measure the quantity of a target sequence

(commonly in real-time). It quantitatively measures starting amounts of DNA, cDNA, or

RNA.quantitative PCR is commonly used to determine whether a DNA sequence is present in a

sample and the number of its copies in the sample. Quantitative PCR has a very high degree of

precision. Quantitative PCR methods use fluorescent dyes, such as Syber Green, EvaGreen

or fluorophore-containing DNA probes, such as TaqMan, to measure the amount of amplified

product in real time. It is also sometimes abbreviated to RT-PCR (real-time PCR) but this

abbreviation should be used only for reverse transcription PCR. qPCR is the appropriate

contractions for quantitative PCR (real-time PCR).

Reverse Transcription PCR (RT-PCR): for amplifying DNA from RNA. Reverse

transcriptase reverse transcribes RNA into cDNA, which is then amplified by PCR. RT-PCR is

widely used in expression profiling, to determine the expression of a gene or to identify the

sequence of an RNA transcript, including transcription start and termination sites. If the

genomic DNA sequence of a gene is known, RT-PCR can be used to map the location

of exons and introns in the gene. The 5' end of a gene (corresponding to the transcription start

site) is typically identified by RACE-PCR (Rapid Amplification of cDNA Ends).

Solid Phase PCR: encompasses multiple meanings, including Polony Amplification (where

PCR colonies are derived in a gel matrix, for example), Bridge PCR[44] (primers are covalently

linked to a solid-support surface), conventional Solid Phase PCR (where Asymmetric PCR is

applied in the presence of solid support bearing primer with sequence matching one of the

aqueous primers) and Enhanced Solid Phase PCR[4 5] (where conventional Solid Phase PCR can

be improved by employing high Tm and nested solid support primer with optional application

of a thermal 'step' to favour solid support priming).

Suicide PCR: typically used in paleogenetics or other studies where avoiding false positives

and ensuring the specificity of the amplified fragment is the highest priority. It was originally

30

described in a study to verify the presence of the microbe Yersinia pestis in dental samples

obtained from 14th Century graves of people supposedly killed by plague during the

medieval Black Death epidemic.[46 ] The method prescribes the use of any primer combination

only once in a PCR (hence the term "suicide"), which should never have been used in any

positive control PCR reaction, and the primers should always target a genomic region never

amplified before in the lab using this or any other set of primers. This ensures that no

contaminating DNA from previous PCR reactions is present in the lab, which could otherwise

generate false positives.

Thermal asymmetric interlaced PCR (TAIL-PCR): for isolation of an unknown

sequence flanking a known sequence. Within the known sequence, TAIL-PCR uses a nested pair

of primers with differing annealing temperatures; a degenerate primer is used to amplify in the

other direction from the unknown sequence.[4 7]

Touchdown PCR (Step-down PCR): a variant of PCR that aims to reduce nonspecific

background by gradually lowering the annealing temperature as PCR cycling progresses. The

annealing temperature at the initial cycles is usually a few degrees (3-5 °C) above the Tm of the

primers used, while at the later cycles, it is a few degrees (3-5 °C) below the primer Tm. The

higher temperatures give greater specificity for primer binding, and the lower temperatures

permit more efficient amplification from the specific products formed during the initial

cycles.[48]

Universal Fast Walking: for genome walking and genetic fingerprinting using a more

specific 'two-sided' PCR than conventional 'one-sided' approaches (using only one gene-specific

primer and one general primer—which can lead to artefactual 'noise')[49] by virtue of a

mechanism involving lariat structure formation. Streamlined derivatives of UFW are LaNe

RAGE (lariat-dependent nested PCR for rapid amplification of genomic DNA ends),[5 0] 5'RACE

LaNe[51] and 3'RACE LaNe.[5 2]

History

Diagrammatic representation of an example primer pair. The use of primers in an in vitro assay to allow DNA

synthesis was a major innovation that allowed the development of PCR.

31

A 1971 paper in the Journal of Molecular Biology by Kleppe and co-workers first described a method using an enzymatic assay to replicate a short DNA template with primers in vitro.[53] However, this early manifestation of the basic PCR principle did not receive much attention, and the invention of the polymerase chain reaction in 1983 is generally credited to Kary Mullis.[54]

"Baby Blue", a 1986 prototype machine for doing PCR

When Mullis developed the PCR in 1983, he was working in Emeryville, California for Cetus Corporation, one of the first biotechnologycompanies. There, he was responsible for synthesizing short chains of DNA. Mullis has written that he conceived of PCR while cruising along the Pacific Coast Highway one night in his car.[55] He was playing in his mind with a new way of analyzing changes (mutations) in DNA when he realized that he had instead invented a method of amplifying any DNA region through repeated cycles of duplication driven by DNA polymerase. In Scientific American, Mullis summarized the procedure: "Beginning with a single molecule of the genetic material DNA, the PCR can generate 100 billion similar molecules in an afternoon. The reaction is easy to execute. It requires no more than a test tube, a few simple reagents, and a source of heat."[56] He was awarded the Nobel Prize in Chemistry in 1993 for his invention,[5] seven years after he and his colleagues at Cetus first put his proposal to practice. However, some controversies have remained about the intellectual and practical contributions of other scientists to Mullis' work, and whether he had been the sole inventor of the PCR principle (see below).

At the core of the PCR method is the use of a suitable DNA polymerase able to withstand the high temperatures of >90 °C (194 °F) required for separation of the two DNA strands in the DNA double helix after each replication cycle. The DNA polymerases initially employed for in vitro experiments presaging PCR were unable to withstand these high temperatures. [3] So the early procedures for DNA replication were very inefficient and time consuming, and required large amounts of DNA polymerase and continuous handling throughout the process.

The discovery in 1976 of Taq polymerase — a DNA polymerase purified from the thermophilic bacterium, Thermus aquaticus, which naturally lives in hot (50 to 80 °C (122 to 176 °F)) environments[12 ] such as hot springs — paved the way for dramatic improvements of the PCR method. The DNA polymerase isolated from T. aquaticus is stable at high temperatures remaining active even after DNA denaturation,[13 ] thus obviating the need to add new DNA polymerase after each cycle.[4] This allowed an automated thermocycler-based process for DNA amplification.

Patent disputes

The PCR technique was patented by Kary Mullis and assigned to Cetus Corporation, where Mullis worked when he invented the technique in 1983. The Taq polymerase enzyme was also covered by patents. There have been several high-profile lawsuits related to the technique, including an unsuccessful lawsuit brought by DuPont. The pharmaceutical company Hoffmann-La Roche purchased the rights to the patents in 1992 and currently holds those that are still protected.

32

Variants of PCR

Basic modifications

Often only a small modification needs to be made to the standard PCR protocol to achieve a desired goal:

Multiplex-PCR uses several pairs of primers annealing to different target sequences.

This permits the simultaneous analysis of multiple targets in a single sample. For example,

in testing for genetic mutations, six or more amplifications might be combined. In the

standard protocol for DNA Fingerprinting, the targets assayed are often amplified in groups

of 3 or 4. Multiplex Ligation-dependent Probe Amplification (or MLPA) permits multiple

targets to be amplified using only a single pair of primers, avoiding the resolution

limitations of multiplex PCR. Multiplex PCR has also been used for analysis

of microsatellites and SNPs.[1 ]

Variations in VNTR lengths in 6 individuals.

Variable Number of Tandem Repeats (VNTR) PCR targets areas of the

genome that exhibit length variation. The analysis of the genotypes of the sample usually

involves sizing of the amplification products by gel electrophoresis. Analysis of smaller

VNTR segments known as Short Tandem Repeats (or STRs) is the basis for DNA

Fingerprinting databases such as CODIS.

Asymmetric PCR preferentially amplifies one strand of the target DNA. It is used in

some sequencing methods and hybridization probing, to generate one DNA strand as

product. Thermocycling is carried out as in PCR, but with a limiting amount or leaving out

one of the primers. When the limiting primer becomes depleted, replication

increases arithmetically through extension of the excess primer. [2] A modification of this

process, named L'inear-After-The-Exponential-PCR (or LATE-PCR), uses a limiting primer

with a higher Melting temperature (Tm) than the excess primer to maintain reaction

efficiency as the limiting primer concentration decreases mid-reaction.[3](Also see Overlap-

extension PCR).

33

Some modifications are needed to perform long PCR. The original Klenow-based PCR

process did not generate products that were larger than about 400 bp. Taq polymerasecan

however amplify targets of up to several thousand bp long.[4] Since then, modified protocols

with Taq enzyme have allowed targets of over 50 kb to be amplified.

Nested PCR is used to increase the specificity of DNA amplification. Two sets of primers

are used in two successive reactions. In the first PCR, one pair of primers is used to

generate DNA products, which may contain products amplified from non-target areas. The

products from the first PCR are then used as template in a second PCR, using one ('hemi-

nesting') or two different primers whose binding sites are located (nested) within the first

set, thus increasing specificity. Nested PCR is often more successful in specifically

amplifying long DNA products than conventional PCR, but it requires more detailed

knowledge of the sequence of the target.

Quantitative PCR is used to measure the specific amount of target DNA (or RNA) in a

sample. By measuring amplification only within the phase of true exponential increase, the

amount of measured product more accurately reflects the initial amount of target. Special

thermal cyclers are used that monitor the amount of product during the

amplification. Quantitative Real-Time PCR (QRT-PCR) methods use fluorescent dyes, such as

Sybr Green, or fluorophore-containing DNA probes, such as TaqMan, to measure the

amount of amplified product as the amplification progresses.

Hot-start PCR is a technique performed manually by heating the reaction components

to the DNA melting temperature (e.g. 95 °C) before adding the polymerase. In this way,

non-specific amplification at lower temperatures is prevented.[6] Alternatively, specialized

reagents inhibit the polymerase's activity at ambient temperature, either by the binding of

an antibody, or by the presence of covalently bound inhibitors that only dissociate after a

high-temperature activation step. 'Hot-start/cold-finish PCR' is achieved with new hybrid

polymerases that are inactive at ambient temperature and are only activated at elevated

temperatures.

In Touchdown PCR, the annealing temperature is gradually decreased in later cycles.

The annealing temperature in the early cycles is usually 3-5 °C above the standard Tmof the

primers used, while in the later cycles it is a similar amount below the Tm. The initial higher

34

annealing temperature leads to greater specificity for primer binding, while the lower

temperatures permit more efficient amplification at the end of the reaction.[7]

Assembly PCR (also known as Polymerase Cycling Assembly or PCA) is the synthesis of

long DNA structures by performing PCR on a pool of long oligonucleotides with short

overlapping segments, to assemble two or more pieces of DNA into one piece. It involves an

initial PCR with primers that have an overlap and a second PCR using the products as the

template that generates the final full-length product. This technique may substitute

for ligation-based assembly.[8]

In Colony PCR, bacterial colonies are screened directly by PCR, for example, the screen

for correct DNA vector constructs. Colonies are sampled with a sterile pipette tip and a

small quantity of cells transferred into a PCR mix. To release the DNA from the cells, the

PCR is either started with an extended time at 95 °C (when standard polymerase is used),

or with a shortened denaturation step at 100 °C and special chimeric DNA polymerase.[9 ]

The Digital polymerase chain reaction simultaneously amplifies thousands of samples, each

in a separate droplet within an emulsion.

Suicide PCR is typically used in paleogenetics or other studies where avoiding false

positives and ensuring the specificity of the amplified fragment is the highest priority. It

was originally described in a study to verify the presence of the microbe Yersinia pestis in

dental samples obtained from 14th Century graves of people supposedly killed by plague

during the medieval Black Death epidemic.[10] The method prescribes the use of any primer

combination only once in a PCR (hence the term "suicide"), which should never have been

used in any positive control PCR reaction, and the primers should always target a genomic

region never amplified before in the lab using this or any other set of primers. This ensures

that no contaminating DNA from previous PCR reactions is present in the lab, which could

otherwise generate false positives.

35

Pretreatments and extensions The basic PCR process can sometimes precede or follow another technique:

RT-PCR (or Reverse Transcription PCR) is used to reverse-transcribe and

amplify RNA to cDNA. PCR is preceded by a reaction using reverse transcriptase, an enzyme

that converts RNA into cDNA. The two reactions may be combined in a tube, with the initial

heating step of PCR being used to inactivate the transcriptase.[4] The Tth polymerase

(described below) has RT activity, and can carry out the entire reaction. RT-PCR is widely

used in expression profiling, which detects the expression of a gene. It can also be used to

obtain sequence of an RNA transcript, which may aid the determination of the transcription

start and termination sites (by RACE-PCR) and facilitate mapping of the location

of exons and introns in a gene sequence.

Ligation-mediated PCR uses small DNA oligonucleotide 'linkers' (or

adaptors) that are first ligated to fragments of the target DNA. PCR primers that anneal

to the linker sequences are then used to amplify the target fragments. This method is

deployed for DNA sequencing, genome walking, and DNA footprinting.[11 ] A related

technique isAmplified fragment length polymorphism, which generates diagnostic fragments

of a genome.

Methylation-specific PCR (MSP) is used to identify patterns of DNA

methylation at cytosine-guanine (CpG) islands in genomic DNA.[12] Target DNA is first

treated with sodium bisulfite, which converts unmethylated cytosine bases to uracil, which

is complementary to adenosine in PCR primers. Two amplifications are then carried out on

the bisulfite-treated DNA: One primer set anneals to DNA with cytosines (corresponding to

methylated cytosine), and the other set anneals to DNA with uracil (corresponding to

unmethylated cytosine). MSP used in quantitative PCR provides quantitative information

about the methylation state of a given CpG island. [13]

Other modifications

Adjustments of the components in PCR is commonly used for optimal performance:

The divalent magnesium ion (Mg++) is required for PCR polymerase activity. Lower

concentrations Mg++ will increase replication fidelity, while higher concentrations will

introduce more mutations. [cita tion need ed]

Denaturants(such as DMSO) can increase amplification specificity by destabilizing non-

specific primer binding. Other chemicals, such as glycerol, are stabilizers for the activity of

36

the polymerase during amplification. Detergents (such as Triton X-100) can prevent

polymerase stick to itself or to the walls of the reaction tube.

DNA polymerases occasionally incorporate mismatch bases into the extending strand. High-

fidelity PCR employs enzymes with 3'-5' exonuclease activity that decreases this rate of mis-

incorporation. Examples of enzymes with proofreading activity include Pfu; adjustments of

the Mg++ and dNTP concentrations may help maximize the number of products that exactly

match the original target DNA.[c itation needed]

COLD-PCR (co-amplification at lower denaturation temperature-PCR) is a modified

Polymerase Chain Reaction (PCR) protocol that enriches variant alleles from a mixture of

wildtype and mutation-containing DNA.

Primer modifications Adjustments to the synthetic oligonucleotides used as primers in PCR are a rich source of modification:

Normally PCR primers are chosen from an invariant part of the genome, and might be used

to amplify a polymorphic area between them. In Allele-specific PCR the opposite is done.

At least one of the primers is chosen from a polymorphic area, with the mutations located at

(or near) its 3'-end. Under stringent conditions, a mismatched primer will not initiate

replication, whereas a matched primer will. The appearance of an amplification product

therefore indicates the genotype. (For more information, see SNP genotyping.)

InterSequence-Specific PCR (or ISSR-PCR) is method for DNA fingerprinting that uses

primers selected from segments repeated throughout a genome to produce a unique

fingerprint of amplified product lengths.[14 ] The use of primers from a commonly repeated

segment is called Alu-PCR, and can help amplify sequences adjacent (or between) these

repeats.

Primers can also be designed to be 'degenerate' - able to initiate replication from a large

number of target locations. Whole genome amplification (or WGA) is a group of

procedures that allow amplification to occur at many locations in an unknown genome, and

which may only be available in small quantities. Other techniques use degenerate

primers that are synthesized using multiple nucleotides at particular positions (the

polymerase 'chooses' the correctly matched primers). Also, the primers can be synthesized

with the nucleoside analog inosine, which hybridizes to three of the four normal bases. A

similar technique can force PCR to perform Site-directed mutagenesis. (also seeOverlap

extension polymerase chain reaction)

37

Normally the primers used in PCR are designed to be fully complementary to the target.

However, the polymerase is tolerant to mis-matches away from the 3' end. Tailed-

primers include non-complementary sequences at their 5' ends. A common procedure is

the use of linker-primers, which ultimately place restriction sites at the ends of the PCR

products, facilitating their later insertion into cloning vectors.

An extension of the 'colony-PCR' method (above), is the use of vector primers. Target DNA

fragments (or cDNA) are first inserted into a cloning vector, and a single set of primers are

designed for the areas of the vector flanking the insertion site. Amplification occurs for

whatever DNA has been inserted. [4]

PCR can easily be modified to produce a labeled product for subsequent use as

a hybridization probe. One or both primers might be used in PCR with a radioactive or

fluorescent label already attached, or labels might be added after amplification. These

labeling methods can be combined with 'asymmetric-PCR' (above) to produce effective

hybridization probes.

38

DNA Polymerases There are several DNA polymerases that are used in PCR:

The Klenow fragment, derived from the original DNA Polymerase I from E. coli, was the first

enzyme used in PCR. Because of its lack of stability at high temperature, it needs be

replenished during each cycle, and therefore is not commonly used in PCR.

The bacteriophage T4 DNA polymerase was also initially used in PCR. It has a higher fidelity

of replication than the Klenow fragment, but is also destroyed by heat.

The DNA polymerase from Thermus aquaticus (or Taq), was the first thermostable

polymerase used in PCR,[4] and is still the one most commonly used. The enzyme can be

isolated from its native source, or from its cloned gene expressed in E. coli.

The Stoffel fragment is made from a truncated gene for Taq polymerase and expressed in E.

coli. It is lacking 5'-3' exonuclease activity, and may be able to amplify longer targets than

the native enzyme. It is a 61 kDa modified form of recombinant AmpliTaq -

see http://www6.appliedbiosystems.com/support/tutorials/pcropt/[c itation n eeded]

Faststart polymerase is a variant of Taq polymerase that requires strong heat activation,

thereby avoiding non-specific amplification due to polymerase activity at low temperature

(see hot-start PCR above).

Pfu DNA polymerase, isolated from the archean Pyrococcus furiosus, has proofreading

activity, and a 5-fold decrease in the error rate of replication compared to Taq. [15]Since

errors increase as PCR progresses, Pfu is the preferred polymerase when products are to be

individually cloned for sequencing or expression.

Vent polymerase is an extremely thermostable DNA polymerase isolated from Thermococcus

litoralis.

Pwo DNA polymerase is a lesser used enzyme obtained from Pyrococcus woesei from which

it takes its name.

Tth polymerase is a thermostable polymerase from Thermus thermophilus. It has reverse

transcriptase activity in the presence of Mn2+ ions, allowing PCR amplification from RNA

targets.

39

Mechanism modifications

Sometimes even the basic mechanism of PCR can be modified:

Inverse PCR.

Unlike normal PCR, Inverse PCR allows amplification and sequencing of DNA that

surrounds a known sequence. It involves initially subjecting the target DNA to a series

of restriction enzyme digestions, and then circularizing the resulting fragments by self

ligation. Primers are designed to be extended outward from the known segment,

resulting in amplification of the rest of the circle. This is especially useful in identifying

sequences to either side of various genomic inserts.[16 ]

Similarly, Thermal Asymmetric InterLaced PCR (or TAIL-PCR) is used to isolate

unknown sequences flanking a known area of the genome. Within the known sequence,

TAIL-PCR uses a nested pair of primers with differing annealing temperatures. A

'degenerate' primer is used to amplify in the other direction from the unknown

sequence.[17]

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Isothermal amplification methods

Some DNA amplification protocols have been developed that may be used alternatively to PCR:

Helicase-dependent amplification is similar to traditional PCR, but uses a constant

temperature rather than cycling through denaturation and annealing/extension

steps. DNA Helicase, an enzyme that unwinds DNA, is used in place of thermal

denaturation.[1 8]

PAN-AC also uses isothermal conditions for amplification, and may be used to analyze

living cells.[19][2 0]

Nicking Enzyme Amplification Reaction referred to as NEAR, is isothermal, replicating

DNA at a constant temperature using a polymerase and nicking enzyme.

Recombinase Polymerase Amplification (RPA).[21] The method uses a recombinase to

specifically pair primers with double-stranded DNA on the basis of homology, thus

directing DNA synthesis from defined DNA sequences present in the sample. Presence

of the target sequence initiates DNA amplification, and no thermal or chemical melting

of DNA is required. The reaction progresses rapidly and results in specific DNA

amplification from just a few target copies to detectable levels typically within 5–10

minutes. The entire reaction system is stable as a dried formulation and does not need

refrigeration. RPA can be used to replace PCR (Polymerase Chain Reaction) in a variety

of laboratory applications and users can design their own assays.[2 2]

41

Gel electrophoresis Gel electrophoresis

Gel electrophoresis apparatus – An agarose gel is placed in this

buffer-filled box and an electrical field is applied via the power

supply to the rear. The negative terminal is at the far end (black

wire), so DNA migrates toward the anode (red wire).

Gel electrophoresis is a method for separation and analysis of macromolecules (DNA, RNA and proteins) and their fragments, based on their size and charge. It is used in clinical chemistry to separate proteins by charge and/or size (IEF agarose, essentially size independent) and in biochemistry and molecular biology to separate a mixed population of DNA and RNA fragments by length, to estimate the size of DNA and RNA fragments or to separate proteins by charge.[1]

Nucleic acid molecules are separated by applying an electric field to move the negatively charged molecules through a matrix ofagarose or other substances. Shorter molecules move faster and migrate farther than longer ones because shorter molecules migrate more easily through the pores of the gel. This phenomenon is called sieving. [2]

Proteins are separated by charge in agarose because the pores of the gel are too large to sieve proteins.

Gel electrophoresis can also be used for separation of nanoparticles.

Gel electrophoresis uses a gel as an anticonvective medium and/or sieving medium during electrophoresis, the movement of a charged particle in an electrical field. Gels suppress the thermal convection caused by application of the electric field, and can also act as a sieving medium, retarding the passage of molecules; gels can also simply serve to maintain the finished separation,

42

so that a post electrophoresis stain can be applied.[3] DNA Gel electrophoresis is usually performed for analytical purposes, often after amplification of DNA via PCR, but may be used as a preparative technique prior to use of other methods such as mass spectrometry,RFLP, PCR, cloning, DNA sequencing, or Southern blotting for further characterization.

Physical basis In simple terms, electrophoresis is a process which enables the sorting of molecules based on size. Using an electric field, molecules (such as DNA) can be made to move through a gel made of agar or polyacrylamide. The electric field consists of a negative charge at one end which pushes the molecules through the gel, and a positive charge at the other end that pulls the molecules through the gel. The molecules being sorted are dispensed into a well in the gel material. The gel is placed in an electrophoresis chamber, which is then connected to a power source. When the electric current is applied, the larger molecules move more slowly through the gel while the smaller molecules move faster. The different sized molecules form distinct bands on the gel.[c itation needed]

The term "gel" in this instance refers to the matrix used to contain, then separate the target molecules. In most cases, the gel is a crosslinked polymer whose composition and porosity is chosen based on the specific weight and composition of the target to be analyzed. When separating proteins or small nucleic acids (DNA, RNA, or oligonucleotides) the gel is usually composed of different concentrations ofacrylamide and a cross-linker, producing different sized mesh networks of polyacrylamide. When separating larger nucleic acids (greater than a few hundred bases), the preferred matrix is purified agarose. In both cases, the gel forms a solid, yet porous matrix. Acrylamide, in contrast to polyacrylamide, is a neurotoxin and must be handled using appropriate safety precautions to avoid poisoning. Agarose is composed of long unbranched chains of uncharged carbohydrate without cross links resulting in a gel with large pores allowing for the separation of macromolecules and macromolecular complexes.[c itation n eeded]

"Electrophoresis" refers to the electromotive force (EMF) that is used to move the molecules through the gel matrix. By placing the molecules in wells in the gel and applying an electric field, the molecules will move through the matrix at different rates, determined largely by their mass when the charge to mass ratio (Z) of all species is uniform. However when charges are not all uniform then, the electrical field generated by the electrophoresis procedure will affect the species that have different charges and therefore will attract the species according to their charges being the opposite. Species that are positively charged (cations) will migrate towards the cathode which is negatively charged. If the species are negatively charged (anions) they will migrate towards the positively charged anode.[4]

If several samples have been loaded into adjacent wells in the gel, they will run parallel in individual lanes. Depending on the number of different molecules, each lane shows separation of the components from the original mixture as one or more distinct bands, one band per component. Incomplete separation of the components can lead to overlapping bands, or to indistinguishable smears representing multiple unresolved components.[c itation needed] Bands in different lanes that end up at the same distance from the top contain molecules that passed through the gel with the same speed, which usually means they are approximately the same size. There are molecular weight size markersavailable that contain a mixture of molecules of known sizes. If such a marker was run on one lane in the gel parallel to the unknown samples, the bands observed can be compared to those of the unknown in order to determine their size. The distance a band travels is approximately inversely proportional to the logarithm of the size of the molecule.[c itation needed]

There are limits to electrophoretic techniques. Since passing current through a gel causes heating, gels may melt during electrophoresis. Electrophoresis is performed in buffer solutions to reduce pH

43

changes due to the electric field, which is important because the charge of DNA and RNA depends on pH, but running for too long can exhaust the buffering capacity of the solution. Further, different preparations of genetic material may not migrate consistently with each other, for morphological or other reasons.

Types of gel The types of gel most typically used are agarose and polyacrylamide gels. Each type of gel is well-suited to different types and sizes of analyte. Polyacrylamide gels are usually used for proteins, and have very high resolving power for small fragments of DNA (5-500 bp). Agarose gels on the other hand have lower resolving power for DNA but have greater range of separation, and are therefore used for DNA fragments of usually 50-20,000 bp in size, but resolution of over 6 Mb is possible with pulsed field gel electrophoresis(PFGE).[5] Polyacrylamide gels are run in a vertical configuration while agarose gels are typically run horizontally in a submarine mode. They also differ in their casting methodology, as agarose sets thermally, while polyacrylamide forms in a chemical polymerization reaction.

Agarose

Agarose gels are made from the natural polysaccharide polymers extracted from seaweed. Agarose gels are easily cast and handled compared to other matrices, because the gel setting is a physical rather than chemical change. Samples are also easily recovered. After the experiment is finished, the resulting gel can be stored in a plastic bag in a refrigerator.

Agarose gels do not have a uniform pore size, but are optimal for electrophoresis of proteins that are larger than 200 kDa.[6] Agarose gel electrophoresis can also be used for the separation of DNA fragments ranging from 50 base pair to several megabases (millions of bases), the largest of which require specialized apparatus. The distance between DNA bands of different lengths is influenced by the percent agarose in the gel, with higher percentages requiring longer run times, sometimes days. Instead high percentage agarose gels should be run with a pulsed field electrophoresis (PFE), or field inversion electrophoresis.

"Most agarose gels are made with between 0.7% (good separation or resolution of large 5–10kb DNA fragments) and 2% (good resolution for small 0.2–1kb fragments) agarose dissolved in electrophoresis buffer. Up to 3% can be used for separating very tiny fragments but a vertical polyacrylamide gel is more appropriate in this case. Low percentage gels are very weak and may break when you try to lift them. High percentage gels are often brittle and do not set evenly. 1% gels are common for many applications."[7 ]

Polyacrylamide

Polyacrylamide gel electrophoresis (PAGE) is used for separating proteins ranging in size from 5 to 2,000 kDa due to the uniform pore size provided by the polyacrylamide gel. Pore size is controlled by modulating the concentrations of acrylamide and bis-acrylamide powder used in creating a gel. Care must be used when creating this type of gel, as acrylamide is a potent neurotoxin in its liquid and powdered forms.

Traditional DNA sequencing techniques such as Maxam-Gilbert or Sanger methods used polyacrylamide gels to separate DNA fragments differing by a single base-pair in length so the sequence could be read. Most modern DNA separation methods now use agarose gels, except for particularly small DNA fragments. It is currently most often used in the field of immunology and protein analysis, often used to separate different proteins or isoforms of the same protein into separate bands. These can be transferred onto anitrocellulose or PVDF membrane to be probed with antibodies and corresponding markers, such as in a western blot.

44

Typically resolving gels are made in 6%, 8%, 10%, 12% or 15%. Stacking gel (5%) is poured on top of the resolving gel and a gel comb (which forms the wells and defines the lanes where proteins, sample buffer and ladders will be placed) is inserted. The percentage chosen depends on the size of the protein that one wishes to identify or probe in the sample. The smaller the known weight, the higher the percentage that should be used. Changes on the buffer system of the gel can help to further resolve proteins of very small sizes. [8]

Starch

Partially hydrolysed potato starch makes for another non-toxic medium for protein electrophoresis. The gels are slightly more opaque than acrylamide or agarose. Non-denatured proteins can be separated according to charge and size. They are visualised using Napthal Black or Amido Black staining. Typical starch gel concentrations are 5% to 10%.[9 ][10 ][11]

Gel conditions Denaturing

Denaturing gels are run under conditions that disrupt the natural structure of the analyte, causing it to unfold into a linear chain. Thus, the mobility of each macromolecule depends only on its linear length and its mass-to-charge ratio. Thus, the secondary, tertiary, and quaternary levels of biomolecular structure are disrupted, leaving only the primary structure to be analyzed.

Nucleic acids are often denatured by including urea in the buffer, while proteins are denatured using sodium dodecyl sulfate, usually as part of the SDS-PAGE process. For full denaturation of proteins, it is also necessary to reduce the covalent disulfide bonds that stabilize their tertiary and quaternary structure, a method called reducing PAGE. Reducing conditions are usually maintained by the addition of beta-mercaptoethanol or dithiothreitol. For general analysis of protein samples, reducing PAGE is the most common form of protein electrophoresis.

Denaturing conditions are necessary for proper estimation of molecular weight of RNA. RNA is able to form more intramolecular interactions than DNA which may result in change of its electrophoretic mobility. Urea,DMSO and glyoxal are the most often used denaturing agents to disrupt RNA structure. Originally, highly toxicmethylmercury hydroxide was often used in denaturing RNA electrophoresis,[12] but it may be method of choice for some samples.[13]

Denaturing gel electrophoresis is used in the DNA and RNA banding pattern-based methods DGGE (denaturing gradient gel electrophoresis),[14] TGGE (temperature gradient gel electrophoresis), and TTGE (temporal temperature gradient electrophoresis).[15]

Native

Native gels are run in non-denaturing conditions, so that the analyte's natural structure is maintained. This allows the physical size of the folded or assembled complex to affect the mobility, allowing for analysis of all four levels of the biomolecular structure. For biological samples, detergents are used only to the extent that they are necessary to lyse lipid membranes in the cell. Complexes remain—for the most part—associated and folded as they would be in the cell. One downside, however, is that complexes may not separate cleanly or predictably, as it is difficult to predict how the molecule's shape and size will affect its mobility.

45

Unlike denaturing methods, native gel electrophoresis does not use a charged denaturing agent. The molecules being separated (usuallyproteins or nucleic acids) therefore differ not only in molecular mass and intrinsic charge, but also the cross-sectional area, and thus experience different electrophoretic forces dependent on the shape of the overall structure. For proteins, since they remain in the native state they may be visualised not only by general protein staining reagents but also by specific enzyme-linked staining.

Native gel electrophoresis is typically used in proteomics and metallomics.[17] However, native PAGE is also used to scan genes (DNA) for unknown mutations as in Single-strand conformation polymorphism.

Buffers

Buffers in gel electrophoresis are used to provide ions that carry a current and to maintain the pH at a relatively constant value. There are a number of buffers used for electrophoresis. The most common being, for nucleic acids Tris/Acetate/EDTA (TAE), Tris/Borate/EDTA(TBE). Many other buffers have been proposed, e.g. lithium borate, which is almost never used, based on Pubmed citations (LB), iso electric histidine, pK matched goods buffers, etc.; in most cases the purported rationale is lower current (less heat) and or matched ion mobilities, which leads to longer buffer life. Borate is problematic; Borate can polymerize, and/or interact with cis diols such as those found in RNA. TAE has the lowest buffering capacity but provides the best resolution for larger DNA. This means a lower voltage and more time, but a better product. LB is relatively new and is ineffective in resolving fragments larger than 5 kbp; However, with its low conductivity, a much higher voltage could be used (up to 35 V/cm), which means a shorter analysis time for routine electrophoresis. As low as one base pair size difference could be resolved in 3% agarose gel with an extremely low conductivity medium (1 mM Lithium borate).[18]

Most SDS-PAGE protein separations are performed using a "discontinuous" (or DISC) buffer system that significantly enhances the sharpness of the bands within the gel. During electrophoresis in a discontinuous gel system, an ion gradient is formed in the early stage of electrophoresis that causes all of the proteins to focus into a single sharp band in a process called isotachophoresis. Separation of the proteins by size is achieved in the lower, "resolving" region of the gel. The resolving gel typically has a much smaller pore size, which leads to a sieving effect that now determines the electrophoretic mobility of the proteins.

Visualization

After the electrophoresis is complete, the molecules in the gel can be stained to make them visible. DNA may be visualized using ethidium bromide which, when intercalated into DNA, fluoresce under ultraviolet light, while protein may be visualised using silver stain or Coomassie Brilliant Blue dye. Other methods may also be used to visualize the separation of the mixture's components on the gel. If the molecules to be separated contain radioactivity, for example in a DNA sequencing gel, an autoradiogram can be recorded of the gel. Photographs can be taken of gels, often using a Gel Doc system.

Downstream processing

After separation, an additional separation method may then be used, such as isoelectric focusing or SDS-PAGE. The gel will then be physically cut, and the protein complexes extracted from each portion separately. Each extract may then be analysed, such as by peptide mass

46

fingerprinting or de novo peptide sequencing after in-gel digestion. This can provide a great deal of information about the identities of the proteins in a complex.

Applications

Estimation of the size of DNA molecules following restriction enzyme digestion, e.g.

in restriction mapping of cloned DNA.

Analysis of PCR products, e.g. in molecular genetic diagnosis or genetic fingerprinting

Separation of restricted genomic DNA prior to Southern transfer, or of RNA prior

to Northern transfer.

Gel electrophoresis is used in forensics, molecular biology, genetics, microbiology and biochemistry. The results can be analyzed quantitatively by visualizing the gel with UV light and a gel imaging device. The image is recorded with a computer operated camera, and the intensity of the band or spot of interest is measured and compared against standard or markers loaded on the same gel. The measurement and analysis are mostly done with specialized software.

Depending on the type of analysis being performed, other techniques are often implemented in conjunction with the results of gel electrophoresis, providing a wide range of field-specific applications.

Nucleic acids

An agarose gel of a PCR product compared to a DNA ladder.

In the case of nucleic acids, the direction of migration, from negative to positive electrodes, is due to the naturally occurring negative charge carried by their sugar-phosphate backbone.[19]

Double-stranded DNA fragments naturally behave as long rods, so their migration through the gel is relative to their size or, for cyclic fragments, their radius of gyration. Circular DNA such as plasmids, however, may show multiple bands, the speed of migration may depend on whether it is relaxed or supercoiled. Single-stranded DNA or RNA tend to fold up into molecules with complex shapes and migrate through the gel in a complicated manner based on their tertiary structure. Therefore,

47

agents that disrupt the hydrogen bonds, such as sodium hydroxide or formamide, are used to denature the nucleic acids and cause them to behave as long rods again. [20]

Gel electrophoresis of large DNA or RNA is usually done by agarose gel electrophoresis. See the "Chain termination method" page for an example of a polyacrylamide DNA sequencing gel. Characterization through ligand interaction of nucleic acids or fragments may be performed by mobility shift affinity electrophoresis.

Electrophoresis of RNA samples can be used to check for genomic DNA contamination and also for RNA degradation. RNA from eukaryotic organisms shows distinct bands of 28s and 18s rRNA, the 28s band being approximately twice as intense as the 18s band. Degraded RNA has less sharply defined bands, has a smeared appearance, and intensity ratio is less than 2:1.

Proteins

SDS-PAGE autoradiography – The indicated proteins are present in different concentrations in the two samples.

Proteins, unlike nucleic acids, can have varying charges and complex shapes, therefore they may not migrate into the polyacrylamide gel at similar rates, or at all, when placing a negative to positive EMF on the sample. Proteins therefore, are usually denatured in the presence of a detergent such as sodium dodecyl sulfate (SDS) that coats the proteins with a negative charge.[3] Generally, the amount of SDS bound is relative to the size of the protein (usually 1.4g SDS per gram of protein), so that the resulting denatured proteins have an overall negative charge, and all the proteins have a similar charge to mass ratio. Since denatured proteins act like long rods instead of having a complex tertiary shape, the rate at which the resulting SDS coated proteins migrate in the gel is relative only to its size and not its charge or shape.[3 ]

Proteins are usually analyzed by sodium dodecyl sulfate polyacrylamide gel electrophoresis (SDS-PAGE), by native gel electrophoresis, by quantitative preparative native continuous polyacrylamide gel electrophoresis (QPNC-PAGE), or by 2-D electrophoresis.

Characterization through ligand interaction may be performed by electroblotting or by affinity electrophoresis in agarose or by capillary electrophoresis as for estimation of binding constants and determination of structural features like glycan content through lectin binding.

48

History

1930s – first reports of the use of sucrose for gel electrophoresis

1955 – introduction of starch gels, mediocre separation

1959 – introduction of acrylamide gels; disc electrophoresis (Ornstein and Davis); accurate

control of parameters such as pore size and stability; and (Raymond and Weintraub)

1966 – agar gels[21 ]

1969 – introduction of denaturing agents especially SDS separation of protein subunit

(Weber and Osborn)[22]

1970 – Laemmli separated 28 components of T4 phage using a stacking gel and SDS

1972 – agarose gels with ethidium bromide stain[23]

1975 – 2-dimensional gels (O’Farrell); isoelectric focusing then SDS gel electrophoresis

1977 – sequencing gels

1983 – pulsed field gel electrophoresis enables separation of large DNA molecules

1983 – introduction of capillary electrophoresis

2004 – standardized time of polymerization of PAGE gels enables clean and predictable

separation of native proteins[24 ]

A 1959 book on electrophoresis by Milan Bier cites references from the 1800s.[2 5] However, Oliver Smithies made significant contributions. Bier states: "The method of Smithies ... is finding wide application because of its unique separatory power." Taken in context, Bier clearly implies that Smithies' method is an improvement.

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Macrophage Macrophage

A macrophage of a mouse stretching its "arms" (pseudopodia) to

engulf two particles, possiblypathogens. Trypan Blue Exclusion.

Macrophages (Greek: big eaters, from makros "large" + phagein "eat"; abbr. MΦ), are a type of white blood cell that engulfs and digests cellular debris, foreign substances, microbes, cancer cells, and anything else that does not have the types of proteins specific to the surface of healthy body cells on its surface[1] in a process called phagocytosis. Macrophages were first discovered by Élie Metchnikoff, a Russian bacteriologist, in 1884. [2] They are found in essentially all tissues,[3] where they patrol for potential pathogensby amoeboid movement. They play a critical role in non-specific defense (innate immunity), and also help initiate specific defense mechanisms (adaptive immunity) by recruiting other immune cells such as lymphocytes. In humans, dysfunctional macrophages cause severe diseases such as chronic granulomatous disease that result in frequent infections.

Beyond increasing inflammation and stimulating the immune system, macrophages also play an important anti-inflammatory role and can decrease immune reactions through the release of cytokines. Macrophages that encourage inflammation are called M1 macrophages, whereas those that decrease inflammation and encourage tissue repair are called M2 macrophages. [4] This difference is reflected in their metabolism, where macrophages have the unique ability to metabolize one amino acid, arginine, to either a "killer" molecule (nitric oxide) or a "repair" molecule (ornithine).

Human macrophages are about 21 micrometres (0.00083 in) in diameter[5] and are produced by the differentiation of monocytes in tissues. They can be identified using flow cytometry or immunohistochemical staining by their specific expression of proteins such asCD14, CD40, CD11b, CD64, F4/80 (mice)/EMR1 (human), lysozyme M, MAC-1/MAC-3 and CD68.[6 ]

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Life cycle

When a monocyte enters damaged tissue through the endothelium of a blood vessel, a process known as the leukocyte extravasation, it undergoes a series of changes to become a macrophage. Monocytes are attracted to a damaged site by chemical substances through chemotaxis, triggered by a range of stimuli including damaged cells, pathogens and cytokines released by macrophages already at the site. At some sites such as the testis, macrophages have been shown to populate the organ through proliferation. Unlike short-lived neutrophils, macrophages survive longer in the body up to a maximum of several months.

Function

Phagocytosis Macrophages are highly specialized in removal of dying or dead cells and cellular debris. This role is important in chronic inflammation, as the early stages of inflammation are dominated by neutrophil granulocytes, which are ingested by macrophages if they come of age (see CD31 for a description of this process).[7]

The neutrophils are at first attracted to a site, where they proliferate, before they are phagocytized by the macrophages.[7 ] When at the site, the first wave of neutrophils, after the process of aging and after the first 48 hours, stimulate the appearance of the macrophages whereby these macrophages will then ingest the aged neutrophils.[7]

The removal of dying cells is, to a greater extent, handled by fixed macrophages, which will stay at strategic locations such as the lungs, liver, neural tissue, bone, spleen and connective tissue, ingesting foreign materials such as pathogens and recruiting additional macrophages if needed.

When a macrophage ingests a pathogen, the pathogen becomes trapped in a phagosome, which then fuses with a lysosome. Within the phagolysosome, enzymes and toxic peroxides digest the pathogen. However, some bacteria, such as Mycobacterium tuberculosis, have become resistant to these methods of digestion. Typhoidal Salmonellae too induce their own phagocytosis by host macrophages in vivo, and inhibit digestion by lysosomal action, thereby use macrophages to replicate and cause macrophage apoptosis.[8]Macrophages can digest more than 100 bacteria before they finally die due to their own digestive compounds.

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Role in adaptive immunity Macrophages are versatile cells that play many roles. As scavengers, they rid the body of worn-out cells and other debris. Along with dendritic cells, they are foremost among the cells that present antigens, a crucial role in initiating an immune response. As secretory cells, monocytes and macrophages are vital to the regulation of immune responses and the development of inflammation; they produce a wide array of powerful chemical substances (monokines) including enzymes, complement proteins, and regulatory factors such as interleukin-1. At the same time, they carry receptors for lymphokines that allow them to be "activated" into single-minded pursuit of microbes and tumour cells.

After digesting a pathogen, a macrophage will present the antigen (a molecule, most often a protein found on the surface of the pathogen and used by the immune system for identification) of the pathogen to the corresponding helper T cell. The presentation is done by integrating it into the cell membrane and displaying it attached to an MHC class II molecule, indicating to other white blood cells that the macrophage is not a pathogen, despite having antigens on its surface.

Eventually, the antigen presentation results in the production of antibodies that attach to the antigens of pathogens, making them easier for macrophages to adhere to with their cell membrane and phagocytose. In some cases, pathogens are very resistant to adhesion by the macrophages.

The antigen presentation on the surface of infected macrophages (in the context of MHC class II) in a lymph node stimulates TH1 (type 1 helper T cells) to proliferate (mainly due to IL-12 secretion from the macrophage). When a B-cell in the lymph node recognizes the same unprocessed surface antigen on the bacterium with its surface bound antibody, the antigen is endocytosed and processed. The processed antigen is then presented in MHCII on the surface of the B-cell. T cells that express the T cell receptor which recognizes the antigen-MHCII complex (with co-stimulatory factors- CD40 and CD40L) cause the B-cell to produce antibodies that help opsonisation of the antigen so that the bacteria can be better cleared by phagocytes.

Macrophages provide yet another line of defense against tumor cells and somatic cells infected with fungus or parasites. Once a T cell has recognized its particular antigen on the surface of an aberrant cell, the T cell becomes an activated effector cell, producing chemical mediators known as lymphokines that stimulate macrophages into a more aggressive form.

Macrophage subtypes

Some believe that there are several activated forms of macrophages.[9] In spite of a spectrum of ways to activate macrophages, there are two main groups designated M1 and M2. M1 macrophages, as mentioned earlier (previously referred to as classically or alternatively activated macrophages),[10] M1 "killer" macrophages are activated by LPS andIFN-gamma, and secrete high levels of IL-12 and low levels of IL-10. In contrast, the M2 "repair" designation broadly refers to macrophages that function in constructive processes like wound healing and tissue repair, and those that turn off damaging immune system activation by producing anti-inflammatory cytokines like IL-10. M2 is the phenotype of resident tissue macrophages, and can be further elevated by IL-4. M2 macrophages produce high levels of IL-10, TGF-beta and low levels of IL-12. Tumor-associated macrophages are mainly of the M2 phenotype, and seem to actively promote tumor growth.[11]

Both M1 and M2 macrophages play a role in promotion of atherosclerosis. M1 macrophages promote atherosclerosis by inflammation. M2 macrophages can remove cholesterol from blood vessels, but when the cholesterol is oxidized, the M2 macrophages become apoptotic foam cells contributing to the atheromatous plaque of atherosclerosis.[1 2][13 ]

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Role in muscle regeneration

The first step to understanding the importance of macrophages in muscle repair, growth, and regeneration is that there are two "waves" of macrophages with the onset of damageable muscle use – subpopulations that do and do not directly have an influence on repairing muscle. The initial wave is a phagocytic population that comes along during periods of increased muscle use that are sufficient to cause muscle membrane lysis and membrane inflammation, which can enter and degrade the contents of injured muscle fibers.[1 4][15 ][16 ] These early-invading, phagocytic macrophages reach their highest concentration about 24 hours following the onset of some form of muscle cell injury or reloading.[17] Their concentration rapidly declines after 48 hours.[15] The second group is the non-phagocytic types that are distributed near regenerative fibers. These peak between two and four days and remain elevated for several days during the hopeful muscle rebuilding.[15] The first subpopulation has no direct benefit to repairing muscle, while the second non-phagocytic group does.

It is thought that macrophages release soluble substances that influence the proliferation, differentiation, growth, repair, and regeneration of muscle, but at this time the factor that is produced to mediate these effects is unknown.[17] It is known that macrophages' involvement in promoting tissue repair is not muscle specific; they accumulate in numerous tissues during the healing process phase following injury.[18]

Role in wound healing

Macrophages are essential for wound healing.[19] They replace Polymorphonuclear neutrophils as the predominant cells in the wound by two days after injury.[20] Attracted to the wound site by growth factors released by platelets and other cells, monocytes from the bloodstream enter the area through blood vessel walls.[21 ] Numbers of monocytes in the wound peak one to one and a half days after the injury occurs. Once they are in the wound site, monocytes mature into macrophages. The spleen contains half the body's monocytes in reserve ready to be deployed to injured tissue.[2 2][23 ]

The macrophage's main role is to phagocytize bacteria and damaged tissue, [19] and they also debride damaged tissue by releasing proteases.[2 4] Macrophages also secrete a number of factors such as growth factors and other cytokines, especially during the third and fourth post-wounding days. These factors attract cells involved in the proliferation stage of healing to the area.[25] Macrophages may also restrain the contraction phase.[2 6] Macrophages are stimulated by the low oxygen content of their surroundings to produce factors that induce and speed angiogenesis [27] and they also stimulate cells that reepithelialize the wound, create granulation tissue, and lay down a new extracellular matrix.[28][2 9] By secreting these factors, macrophages contribute to pushing the wound healing process into the next phase.

Role in limb regeneration

Scientists have elucidated that as well as eating up material debris, macrophages are involved in the typical limb regeneration in the salamander.[3 0][31 ] They found that removing the macrophages from a salamander resulted in failure of limb regeneration and a scarring response.[30][31]

Role in iron homeostasis

As described above, macrophages play a key role in removing dying or dead cells and cellular debris. Erythrocytes have a lifespan on average of 120 days and so are constantly being destroyed by macrophages in the spleen and liver. Macrophages will also engulf macromolecules, and so play a key role in the pharmacokinetics of parenteral irons.

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The iron that is released from the haemoglobin is either stored internally in ferritin or is released into the circulation via ferroportin. In cases where systemic iron levels are raised, or where inflammation is present, raised levels of hepcidin act on macrophage ferroportin channels, leading to iron remaining within the macrophages.

Tissue macrophages

Drawing of a macrophage when fixed and stained bygiemsa dye

A majority of macrophages are stationed at strategic points where microbial invasion or accumulation of foreign particles is likely to occur. These cells together as a group are known as the Mononuclear phagocyte system and were previously known as the Reticuloendothelial system. Each type of macrophage, determined by its location, has a specific name:

Cell Name Location

Adipose tissue macrophages Adipose tissue

Monocyte Bone Marrow/Blood

Kupffer cell Liver

Sinus histiocytes Lymph node

Alveolar macrophage(dust cell) Pulmonary alveolus of Lungs

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Tissue macrophage (Histiocyte) leading to Giant cells Connective Tissues

Langerhans cell Skin and Mucosa

Microglia Central Nervous System

Hofbauer cell Placenta

Intraglomerular mesangial cell Kidney

Osteoclasts Bone

Epithelioid cells Granulomas

Red Pulp Macrophage (Sinusoidal lining cells) Red pulp of Spleen

Peritoneal macrophages Peritoneal cavity

Investigations concerning Kupffer cells are hampered because in humans, Kupffer cells are only accessible for immunohistochemical analysis from biopsies or autopsies. From rats and mice, they are difficult to isolate, and after purification, only approximately 5 million cells can be obtained from one mouse.

Macrophages can express paracrine functions within organs that are specific to the function of that organ. In the testis for example, macrophages have been shown to be able to interact with Leydig cells by secreting 25-hydroxycholesterol, an oxysterol that can be converted to testosterone by neighbouring Leydig cells.[32] Also, testicular macrophages may participate in creating an immune privileged environment in the testis, and in mediating infertility during inflammation of the testis.

Macrophages can be classified on basis of the fundamental function and activation. According to this grouping there are classically activated macrophages, wound-healing macrophages (alternatively activated macrophages) and regulatory macrophages (Mregs).[33 ]

Disease Due to their role in phagocytosis, macrophages are involved in many diseases of the immune system. For example, they participate in the formation of granulomas, inflammatory lesions that

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may be caused by a large number of diseases. Some disorders, mostly rare, of ineffective phagocytosis and macrophage function have been described, for example.[c itation needed]

As a host for intracellular pathogens In their role as a phagocytic immune cell macrophages are responsible for engulfing pathogens to destroy them. Some pathogens subvert this process and instead live inside the macrophage. This provides an environment in which the pathogen is hidden from the immune system and allows it to replicate.

Diseases with this type of behaviour include tuberculosis (caused by Mycobacterium tuberculosis) and leishmaniasis (caused by Leishmania species).

Tuberculosis

Once engulfed by a macrophage, the causative agent of tuberculosis, Mycobacterium tuberculosis,[34] avoids cellular defenses and uses the cell to replicate.

Leishmaniasis

Upon phagocytosis by a macrophage, the Leishmania parasite finds itself in a phagocytic vacuole. Under normal circumstances, this phagocytic vacuole would develop into a lysosome and its contents would be digested. Leishmania alter this process and avoid being destroyed; instead, they make a home inside the vacuole.

Chikungunya

Infection of macrophages in joints is associated with local inflammation during and after the acute phase of Chikungunya (caused by CHIKV or Chikungunya Virus).[35]

Others

Adenovirus (most common cause of pink eye) can remain latent in a host macrophage, with continued viral shedding 6–18 months after initial infection.

Brucella spp. can remain latent in a macrophage via inhibition of phagosome–lysosome fusion; causes brucellosis (undulant fever).

Heart disease Macrophages are the predominant cells involved in creating the progressive plaque lesions of atherosclerosis.[36]

HIV infection Macrophages also play a role in Human Immunodeficiency Virus (HIV) infection. Like T cells, macrophages can be infected with HIV, and even become a reservoir of ongoing virus replication throughout the body. HIV can enter the macrophage through binding of gp120 to CD4 and second membrane receptor, CCR5 (a chemokine receptor). Both circulating monocytes and macrophages serve as a reservoir for the virus.[37 ]

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Cancer Macrophages contribute to tumor growth and progression. Attracted to oxygen-starved (hypoxic) and necrotic tumor cells they promote chronic inflammation. Inflammatory compounds such as Tumor necrosis factor (TNF)-alpha released by the macrophages activate the gene switch nuclear factor-kappa B. NF-κB then enters the nucleus of a tumor cell and turns on production of proteins that stop apoptosis and promote cell proliferation and inflammation.[38] Moreover macrophages serve as a source for many pro-angiogenic factors including vascular endothelial factor (VEGF), tumor necrosis factor-alpha (TNF-alpha), granulocyte macrophage colony-stimulating factor (GM-CSF) and IL-1and IL-6 [39] contributing further to the tumor growth. Macrophages have been shown to infiltrate a number of tumors. Their number correlates with poor prognosis in certain cancers including cancers of breast, cervix, bladder and brain.[40] Tumor-associated macrophages (TAMs) are thought to acquire an M2 phenotype, contributing to tumor growth and progression. Recent study findings suggest that by forcing IFN-α expression in tumor-infiltrating macrophages, it is possible to blunt their innate protumoral activity and reprogramme the tumor microenvironment toward more effective dendritic cell activation and immune effector cell cytotoxicity.[41]

Obesity Increased number of pro-inflammatory macrophages within obese adipose tissue contributes to obesity complications including insulin resistance and diabetes type 2.[42 ]

Media

An active J774 macrophage is seen taking up four

conidia in a cooperative manner. The J774 cells were treated with 5 ng/ml interferon-γ one night

before filming with conidia. Observations were made every 30s over a 2.5hr period.

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Electrophoresis .

Illustration of electrophoresis

Illustration of electrophoresis retardation

Electrophoresis is the motion of dispersed particles relative to a fluid under the influence of a spatially uniform electric field.[1][2][3][4 ][5][6] This electrokinetic phenomenon was observed for the first time in 1807 by Ferdinand Frederic Reuss (Moscow State University),[7] who noticed that the application of a constant electric field caused clay particles dispersed inwater to migrate. It is ultimately caused by the presence of a charged interface between the particle surface and the surrounding fluid. It is the basis for a number of analytical techniques used in biochemistry for separating molecules by size, charge, or binding affinity.

Electrophoresis of positively charged particles (cations) is called cataphoresis, while electrophoresis of negatively charged particles (anions) is called anaphoresis. Electrophoresis is a technique used in laboratories in order to separate macromolecules based on size. The technique applies a negative charge so proteins move towards a positive charge. This is used for both DNA

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and RNA analysis. Polyacrylamide gel electrophoresis (PAGE) has a clearer resolution than agarose and is more suitable for quantitative analysis. In this technique DNA foot-printing can identify how proteins bind to DNA. It can be used to separate proteins by size, density and purity. It can also be used for plasmid analysis, which develops our understanding of bacteria becoming resistant to antibiotics.

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Cell culture

Cell culture in a special tissue culture dish

Epithelial cells in culture, stainedfor keratin (red) and DNA (green)

Cell culture is the process by which cells are grown under controlled conditions, generally outside of their natural environment. In practice, the term "cell culture" now refers to the culturing of cells derived from multi-cellular eukaryotes, especially animal cells. However, there are also cultures of plants, fungi, and microbes, including viruses, bacteria and protists. The historical development and methods of cell culture are closely interrelated to those of tissue culture and organ culture.

The laboratory technique of maintaining live cell lines (a population of cells derived from a single cell and containing the same genetic makeup) separated from their original tissue source became more robust in the middle 20th century.[1][2]

History

The 19th-century English physiologist Sydney Ringer developed salt solutions containing the chlorides of sodium, potassium, calcium and magnesium suitable for maintaining the beating of an isolated animal heart outside of the body.[3] In 1885, Wilhelm Roux removed a portion of the medullary plate of an embryonic chicken and maintained it in a warm saline solution for several days, establishing the principle of tissue culture.[4] Ross Granville Harrison, working at Johns Hopkins Medical School and then at Yale University, published results of his experiments from 1907 to 1910, establishing the methodology of tissue culture.[5]

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Cell culture techniques were advanced significantly in the 1940s and 1950s to support research in virology. Growing viruses in cell cultures allowed preparation of purified viruses for the manufacture of vaccines. The injectable polio vaccine developed by Jonas Salk was one of the first products mass-produced using cell culture techniques. This vaccine was made possible by the cell culture research of John Franklin Enders, Thomas Huckle Weller, and Frederick Chapman Robbins, who were awarded a Nobel Prize for their discovery of a method of growing the virus in monkey kidney cell cultures.

Concepts in mammalian cell culture Isolation of cells

Cells can be isolated from tissues for ex vivo culture in several ways. Cells can be easily purified from blood; however, only the white cells are capable of growth in culture. Mononuclear cells can be released from soft tissues by enzymatic digestion with enzymes such as collagenase, trypsin, or pronase, which break down the extracellular matrix. Alternatively, pieces of tissue can be placed in growth media, and the cells that grow out are available for culture. This method is known as explant culture.

Cells that are cultured directly from a subject are known as primary cells. With the exception of some derived from tumors, most primary cell cultures have limited lifespan.

An established or immortalized cell line has acquired the ability to proliferate indefinitely either through random mutation or deliberate modification, such as artificial expression of the telomerase gene. Numerous cell lines are well established as representative of particular cell types.

Maintaining cells in culture

A bottle of DMEM cell culture medium

For the majority of isolated primary cells, they undergo the process of senescence and stop dividing after a certain number of population doublings while generally retaining their viability (described as the Hayflick limit).

Cells are grown and maintained at an appropriate temperature and gas mixture (typically, 37 °C, 5% CO2 for mammalian cells) in a cell incubator. Culture conditions vary widely for each cell type, and variation of conditions for a particular cell type can result in differentphenotypes.

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Aside from temperature and gas mixture, the most commonly varied factor in culture systems is the cell growth medium. Recipes for growth media can vary in pH, glucose concentration, growth factors, and the presence of other nutrients. The growth factors used to supplement media are often derived from the serum of animal blood, such as fetal bovine serum (FBS), bovine calf serum, equine serum, and porcine serum. One complication of these blood-derived ingredients is the potential for contamination of the culture with viruses orprions, particularly in medical biotechnology applications. Current practice is to minimize or eliminate the use of these ingredients wherever possible and use human platelet lysate (hPL). This eliminates the worry of cross-species contamination when using FBS with human cells. hPL has emerged as a safe and reliable alternative as a direct replacement for FBS or other animal serum. In addition,chemically defined media can be used to eliminate any serum trace (human or animal), but this cannot always be accomplished with different cell types. Alternative strategies involve sourcing the animal blood from countries with minimum BSE/TSE risk, such as The United States, Australia and New Zealand,[6] and using purified nutrient concentrates derived from serum in place of whole animal serum for cell culture.[7]

Plating density (number of cells per volume of culture medium) plays a critical role for some cell types. For example, a lower plating density makes granulosa cells exhibit estrogen production, while a higher plating density makes them appear as progesterone-producing theca lutein cells.[8]

Cells can be grown either in suspension or adherent cultures. Some cells naturally live in suspension, without being attached to a surface, such as cells that exist in the bloodstream. There are also cell lines that have been modified to be able to survive in suspension cultures so they can be grown to a higher density than adherent conditions would allow. Adherent cells require a surface, such as tissue culture plastic or microcarrier, which may be coated with extracellular matrix (such as collagen and laminin) components to increase adhesion properties and provide other signals needed for growth and differentiation. Most cells derived from solid tissues are adherent. Another type of adherent culture is organotypic culture, which involves growing cells in a three-dimensional (3-D) environment as opposed to two-dimensional culture dishes. This 3D culture system is biochemically and physiologically more similar to in vivo tissue, but is technically challenging to maintain because of many factors (e.g. diffusion).

Cell line cross-contamination

Cell line cross-contamination can be a problem for scientists working with cultured cells.[9] Studies suggest anywhere from 15–20% of the time, cells used in experiments have been misidentified or contaminated with another cell line. [10][11][12] Problems with cell line cross-contamination have even been detected in lines from the NCI-60 panel, which are used routinely for drug-screening studies.[13][1 4] Major cell line repositories, including the American Type Culture Collection (ATCC), the European Collection of Cell Cultures (ECACC) and the German Collection of Microorganisms and Cell Cultures (DSMZ), have received cell line submissions from researchers that were misidentified by them.[13][1 5]Such contamination poses a problem for the quality of research produced using cell culture lines, and the major repositories are now authenticating all cell line submissions.[16]ATCC uses short tandem repeat (STR) DNA fingerprinting to authenticate its cell lines. [17]

To address this problem of cell line cross-contamination, researchers are encouraged to authenticate their cell lines at an early passage to establish the identity of the cell line. Authentication should be repeated before freezing cell line stocks, every two months during active culturing and before any publication of research data generated using the cell lines. Many methods are used to identify cell lines, including isoenzyme analysis, human lymphocyte antigen (HLA) typing, chromosomal analysis, karyotyping, morphology andSTR analysis.[17 ]

One significant cell-line cross contaminant is the immortal HeLa cell line.

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Other technical issues

As cells generally continue to divide in culture, they generally grow to fill the available area or volume. This can generate several issues:

Nutrient depletion in the growth media

Changes in pH of the growth media

Accumulation of apoptotic/necrotic (dead) cells

Cell-to-cell contact can stimulate cell cycle arrest, causing cells to stop dividing, known

as contact inhibition.

Cell-to-cell contact can stimulate cellular differentiation.

Genetic and epigenetic alterations, with a natural selection of the altered cells potentially

leading to overgrowth of abnormal, culture-adapted cells with decreased differentiation and

increased proliferative capacity.[18]

Manipulation of cultured cells

Among the common manipulations carried out on culture cells are media changes, passaging cells, and transfecting cells. These are generally performed using tissue culture methods that rely on aseptic technique. Aseptic technique aims to avoid contamination with bacteria, yeast, or other cell lines. Manipulations are typically carried out in a biosafety hood or laminar flow cabinet to exclude contaminating micro-organisms. Antibiotics (e.g. penicillin and streptomycin) and antifungals (e.g.amphotericin B) can also be added to the growth media.

As cells undergo metabolic processes, acid is produced and the pH decreases. Often, a pH indicator is added to the medium to measure nutrient depletion.

Media changes

In the case of adherent cultures, the media can be removed directly by aspiration, and then is replaced. Media changes in non-adherent cultures involve centrifuging the culture and resuspending the cells in fresh media.

Passaging cells

Passaging (also known as subculture or splitting cells) involves transferring a small number of cells into a new vessel. Cells can be cultured for a longer time if they are split regularly, as it avoids the senescence associated with prolonged high cell density. Suspension cultures are easily passaged with a small amount of culture containing a few cells diluted in a larger volume of fresh media. For adherent cultures, cells first need to be detached; this is commonly done with a mixture of trypsin-EDTA; however, other enzyme mixes are now available for this purpose. A small number of detached cells can then be used to seed a new culture. Some cell cultures, such as RAW cells are mechanically scraped from the surface of their vessel with rubber scrapers.

Transfection and transduction

Another common method for manipulating cells involves the introduction of foreign DNA by transfection. This is often performed to cause cells to express a protein of interest. More recently, the transfection of RNAi constructs have been realized as a convenient mechanism for

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suppressing the expression of a particular gene/protein. DNA can also be inserted into cells using viruses, in methods referred to as transduction, infection or transformation. Viruses, as parasitic agents, are well suited to introducing DNA into cells, as this is a part of their normal course of reproduction.

Established human cell lines

Cultured HeLa cells have been stained with Hoechst turning theirnuclei blue, and are one of the earliest human cell

lines descended fromHenrietta Lacks, who died of cervical cancer from which these cells originated.

Cell lines that originate with humans have been somewhat controversial in bioethics, as they may outlive their parent organism and later be used in the discovery of lucrative medical treatments. In the pioneering decision in this area, the Supreme Court of California held inMoore v. Regents of the University of California that human patients have no property rights in cell lines derived from organs removed with their consent.[19]

Cell strains

A cell strain is derived either from a primary culture or a cell line by the selection or cloning of cells having specific properties or characteristics which must be defined. Cell strains are cells that have been adapted to culture but, unlike cell lines, have a finite division potential. Non-immortalized cells stop dividing after 40 to 60 population doublings[20] and, after this, they lose their ability to proliferate (a genetically determined event known as senescence).[21]

Applications of cell culture Mass culture of animal cell lines is fundamental to the manufacture of viral vaccines and other products of biotechnology.

Biological products produced by recombinant DNA (rDNA) technology in animal cell cultures include enzymes, synthetic hormones, immunobiologicals (monoclonal antibodies,interleukins, lymphokines), and anticancer agents. Although many simpler proteins can be produced using rDNA in bacterial cultures, more complex proteins that are glycosylated(carbohydrate-modified) currently must be made in animal cells. An important example of such a complex protein is the hormone erythropoietin. The cost of growing mammalian cell cultures is high, so research is underway to produce such complex proteins in insect cells or in higher plants, use of single embryonic cell and somatic embryos as a source for direct gene transfer via particle bombardment, transit gene expression and confocal microscopy observation is one of its applications. It also offers to confirm single cell origin of somatic embryos and the asymmetry of the first cell division, which starts the process.

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Cell culture in two dimensions

Research in tissue engineering, stem cells and molecular biology primarily involves cultures of cells on flat plastic dishes. This technique is known as two-dimensional (2D) cell culture, and was first developed by Wilhelm Roux who, in 1885, removed a portion of the medullary plate of an embryonic chicken and maintained it in warm saline for several days on a flat glass plate. From the advance of polymer technology arose today's standard plastic dish for 2D cell culture, commonly known as the Petri dish. Julius Richard Petri, a German bacteriologist, is generally credited with this invention while working as an assistant to Robert Koch. Various researchers today also utilize culturing laboratory flasks, conicals, and even disposable bags like those used in single-use bioreactors.

Aside from Petri dishes, scientists have long been growing cells within biologically derived matrices such as collagen or fibrin, and more recently, on synthetic hydrogels such as polyacrylamide or PEG. They do this in order to elicit phenotypes that are not expressed on conventionally rigid substrates. There is growing interest in controlling matrix stiffness,[22 ] a concept that has led to discoveries in fields such as:

Stem cell self-renewal

Lineage specification

Cancer cell phenotype

Fibrosis

Hepatocyte function

Mechanosensing

Cell culture in three dimensions

Cell culture in three dimensions has been touted as "Biology's New Dimension".[37] Nevertheless, the practice of cell culture remains based on varying combinations of single or multiple cell structures in 2D.[38] That being said, there is an increase in use of 3D cell cultures in research areas including drug discovery, cancer biology, regenerative medicine and basic life science research.[39] There are a variety of platforms used to facilitate the growth of three-dimensional cellular structures such as nanoparticle facilitated magnetic levitation,[40 ] gel matrices scaffolds, and hanging drop plates.[41]

3D Cell Culturing by Magnetic Levitation

3D Cell Culturing by Magnetic Levitation method (MLM) is the application of growing 3D tissue by inducing cells treated with magnetic nanoparticle assemblies in spatially varying magnetic fields using neodymium magnetic drivers and promoting cell to cell interactions by levitating the cells up to the air/liquid interface of a standard petri dish. The magnetic nanoparticle assemblies consist of magnetic iron oxide nanoparticles, gold nanoparticles, and the polymer polylysine. 3D cell culturing is scalable, with the capability for culturing 500 cells to millions of cells or from single dish to high-throughput low volume systems.

Tissue culture and engineering

Cell culture is a fundamental component of tissue culture and tissue engineering, as it establishes the basics of growing and maintaining cells in vitro. The major application of human cell culture is

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in stem cell industry, where mesenchymal stem cells can be cultured and cryopreserved for future use. Tissue engineering potentially offers dramatic improvements in low cost medical care for hundreds of thousands of patients annually.

Vaccines

Vaccines for polio, measles, mumps, rubella, and chickenpox are currently made in cell cultures. Due to the H5N1 pandemic threat, research into using cell culture for influenza vaccines is being funded by the United States government. Novel ideas in the field include recombinant DNA-based vaccines, such as one made using human adenovirus (a common cold virus) as a vector,[42][43] and novel adjuvants.[44]

Culture of non-mammalian cells Plant cell culture methods

Plant cell cultures are typically grown as cell suspension cultures in a liquid medium or as callus cultures on a solid medium. The culturing of undifferentiated plant cells and calli requires the proper balance of the plant growth hormones auxin and cytokinin.

Insect cell culture

Cells derived from Drosophila melanogaster (most prominently, Schneider 2 cells) can be used for experiments which may be hard to do on live flies or larvae, such asbiochemical studies or studies using siRNA. Cell lines derived from the army worm Spodoptera frugiperda, including Sf9 and Sf21, and from the cabbage looper Trichoplusia ni,High Five cells, are commonly used for expression of recombinant proteins using baculovirus.

Bacterial and yeast culture methods

For bacteria and yeasts, small quantities of cells are usually grown on a solid support that contains nutrients embedded in it, usually a gel such as agar, while large-scale cultures are grown with the cells suspended in a nutrient broth.

Viral culture methods

The culture of viruses requires the culture of cells of mammalian, plant, fungal or bacterial origin as hosts for the growth and replication of the virus. Whole wild type viruses,recombinant viruses or viral products may be generated in cell types other than their natural hosts under the right conditions. Depending on the species of the virus, infection andviral replication may result in host cell lysis and formation of a viral plaque.

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Common cell lines Human cell lines

HeLa

National Cancer Institute's 60 cancer cell lines (NCI60)

ESTDAB database

DU145 (prostate cancer)

Lncap (prostate cancer)

MCF-7 (breast cancer)

MDA-MB-438 (breast cancer)

PC3 (prostate cancer)

T47D (breast cancer)

THP-1 (acute myeloid leukemia)

U87 (glioblastoma)

SHSY5Y Human neuroblastoma cells, cloned from a myeloma

Saos-2 cells (bone cancer)

KBM-7 cells (chronic myelogenous leukemia)

Primate cell lines

Vero (African green monkey Chlorocebus kidney epithelial cell line initiated in 1962) Rat tumor cell lines

GH3 (pituitary tumor)

PC12 (pheochromocytoma) Mouse cell lines

MC3T3 (embryonic calvarium)

Plant cell lines

Tobacco BY-2 cells (kept as cell suspension culture, they are model system of plant cell) Other species cell lines

Zebrafish ZF4 and AB9 cells

Madin-Darby canine kidney (MDCK) epithelial cell line.

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Types of cell cultures Primary cell culture: This is the maintenance of growth of cells dissociated from the

parental tissue (such as kidney or liver) using mechanical or enzymatic methods, in culture medium using suitable glass or plastic containers. The primary cell culture could be of two types depending upon the kind of cells in culture.

o Adherent cells - Cells shown to require attachment for growth are said to be

anchorage dependent cells. The adherent cells are usually derived from tissues of organs such as kidney where they are immobile and embedded in connective tissue.

o Suspension cells - Cells which do not require attachment for growth or do not attach

to the surface of the culture vessels are anchorage independent cells/suspension cells. All suspension cultures are derived from cells of the blood system because these cells are also suspended in plasma in vitro e.g. lymphocytes.

Secondary cell cultures: When a primary culture is sub-cultured, it becomes known as

secondary culture or cell line. Subculture (or passage) refers to the transfer of cells from one culture vessel to another culture vessel. This is periodically required to provide fresh nutrients and growing space for continuously growing cell lines. The process involves removing the growth media and disassociating the adhered cells (usually enzymatically). Such cultures may be called secondary cultures.

Cell Line: A cell line or cell strain may be finite or continuous depending upon whether it has limited culture life span or it is immortal in culture. On the basis of the life span of culture, the cell lines are categorized into two types:

o Finite cell lines - The cell lines which have a limited life span and go through a

limited number of cell generations (usually 20-80 population doublings) are known as finite cell lines. These cell lines exhibit the property of contact inhibition, density limitation and anchorage dependence. The growth rate is slow and doubling time is around 24-96 hours.

o Continuous cell lines - Cell lines transformed under laboratory conditions or in vitro culture conditions give rise to continuous cell lines. These cell lines show the property of ploidy (aneupliody or heteroploidy), absence of contact inhibition and anchorage dependence. They grow either in a monolayer or in suspension (see below). The growth rate is rapid and doubling time can be 12-24 hours.

Monolayer cultures - When the bottom of the culture vessel is covered

with a continuous layer of cells, usually one cell in thickness, they are referred to as monolayer cultures.

Suspension cultures - Majority of continuous cell lines grow as

monolayers. Some of the cells which are non-adhesive e.g. cells of leukemia or certain cells which can be mechanically kept in suspension, can be propagated in suspension.

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Basic cell culture techniques In the laboratory cells are generally maintained in a suitable culture medium either solid or liquid, but in either case one providing an environment suitable for their growth and multiplication. Furthermore, cultures of cells are normally supplied as pure cultures, containing only a single strain. This is in contrast to natural environments where a diversity of microorganisms is usually present. Cultures are usually supplied in conical flasks, universal bottles or Petri dishes, the cotton wool plugs, screw caps or covers respectively serving to keep the cultures free of contamination from airborne bacteria or fungal spores. In order to maintain pure cultures, all glassware must be sterilised before use and aseptic techniques (sometimes known as sterile techniques) should be observed throughout, as described in the following notes.

Transfers of cells from one vessel to another should be carried out as rapidly as possible, though without unnecessary haste and without the need of a partner to hold anything.

Plugs and caps must be held in the fingers when temporarily removed from culture vessels and must be set down on the bench.

The mouths of culture tubes or bottles should be flamed after removing plugs and caps and again before they are replaced.

Work in a laminar flow or close to a Bunsen flame to ensure that airbo rne contaminants are carried upwards.

As the use of microorganisms in producing valuable materials has expanded, attention has turned to using plant and animal cells and tissues in the same way. Although they are more difficult to grow, some progress has been made. One area of cell culture already making a valuable contribution is the manufacture of antibodies, in particular single cell antibodies (monoclonal antibodies).

As β-lymphocytes which produce antibodies are hard to grow outside the body, the production of monoclonal antibodies in any quantity was impossible until recently, when the desired β-lymphocytes were fused with cancer cells. Cancer cells divide rapidly and these new hybridoma cells were found to do so as well, giving a continual source of cells producing a monoclonal antibody.

Monoclonal antibodies have many uses, including estimating the quantity of a chemical in a mixture, a technique called immunoassay. They are therefore used to detect the hormones present in urine (pregnancy testing kits), in detecting drugs in urine (e.g for athletes) and in detecting the human immune deficiency virus (AIDS test for HIV).

Check out the youtube video describing the basics of cell culture techniques.

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Growth medium

An agar plate -- an example of a bacterial growth medium. Specifically, it is a streak plate; the orange lines and dots are formed by bacterial colonies.

A growth medium or culture medium is a liquid or gel designed to support the growth of microorganisms or cells,[1] or small plantslike the moss Physcomitrella patens.[2] There are different types of media for growing different types of cells.[3]

There are two major types of growth media: those used for cell culture, which use specific cell types derived from plants or animals, and microbiological culture, which are used for growing microorganisms, such as bacteria or yeast. The most common growth media for microorganisms are nutrient broths and agar plates; specialized media are sometimes required for microorganism and cell culture growth.[1] Some organisms, termed fastidious organisms, require specialized environments due to complex nutritional requirements.Viruses, for example, are obligate intracellular parasites and require a growth medium containing living cells.

Types of growth media The most common growth media for microorganisms are nutrient broths (liquid nutrient medium) or LB medium (Lysogeny Broth). Liquid media are often mixed with agar and poured via sterile media dispenser into Petri dishes to solidify. These agar plates provide a solid medium on which microbes may be cultured. They remain solid, as very few bacteria are able to decompose agar (the exception being some species in the following genera: Cytophaga, Flavobacterium, Bacillus, Pseudomonas, and Alcaligenes). Bacteria grown in liquid cultures often form colloidal suspensions.[4 ][5]

The difference between growth media used for cell culture and those used for microbiological culture is that cells derived from whole organisms and grown in culture often cannot grow without the addition of, for instance, hormones or growth factors which usually occur in vivo.[6] In the case of animal cells, this difficulty is often addressed by the addition ofblood serum or a synthetic serum replacement to the medium. In the case of microorganisms, there are no such limitations, as they are often unicellular organisms. One other major difference is that animal cells in culture are often grown on a flat surface to which they attach, and the medium is provided in a liquid form, which covers the cells. In contrast, bacteria such as Escherichia coli may be grown on solid media or in liquid media.

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An important distinction between growth media types is that of defined versus undefined media.[1] A defined medium will have known quantities of all ingredients. For microorganisms, they consist of providing trace elements and vitamins required by the microbe and especially a defined carbon source and nitrogen source. Glucose or glycerolare often used as carbon sources, and ammonium salts or nitrates as inorganic nitrogen sources. An undefined medium has some complex ingredients, such as yeast extract or casein hydrolysate, which consist of a mixture of many, many chemical species in unknown proportions. Undefined media are sometimes chosen based on price and sometimes by necessity - some microorganisms have never been cultured on defined media.

A good example of a growth medium is the wort used to make beer. The wort contains all the nutrients required for yeast growth, and under anaerobic conditions, alcohol is produced. When the fermentation process is complete, the combination of medium and dormant microbes, now beer, is ready for consumption.

Nutrient media

Nutrient media contain all the elements that most bacteria need for growth and are non-selective, so they are used for the general cultivation and maintenance of bacteria kept in laboratory culture collections.

Physcomitrella patens plants growing axenically on agar plates (Petri dish, 9 cm diameter).

An undefined medium (also known as a basal or complex medium) is a medium that contains:

a carbon source such as glucose for bacterial growth

water

various salts needed for bacterial growth

a source of amino acids and nitrogen (e.g., beef, yeast extract)

o This is an undefined medium because the amino acid source contains a variety of

compounds with the exact composition being unknown.

A defined medium (also known as chemically defined medium or synthetic medium) is a medium in which

all the chemicals used are known

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no yeast, animal or plant tissue is present

Some examples of nutrient media include:

Plate count agar

Nutrient agar

Trypticase soy agar

Minimal media Minimal media are those that contain the minimum nutrients possible for colony growth, generally without the presence of amino acids, and are often used by microbiologists and geneticists to grow "wild type" microorganisms. Minimal media can also be used to select for or against recombinants or exconjugants.

Minimal medium typically contains:

a carbon source for bacterial growth, which may be a sugar such as glucose, or a less

energy-rich source like succinate

various salts, which may vary among bacteria species and growing conditions; these

generally provide essential elements such as magnesium, nitrogen, phosphorus,

andsulfur to allow the bacteria to synthesize protein and nucleic acid

water

Supplementary minimal media are a type of minimal media that also contains a single selected agent, usually an amino acid or a sugar. This supplementation allows for the culturing of specific lines of auxotrophic recombinants.

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Selective media Selective media are used for the growth of only selected microorganisms. For example, if a microorganism is resistant to a certainantibiotic, such as ampicillin or tetracycline, then that antibiotic can be added to the medium in order to prevent other cells, which do not possess the resistance, from growing. Media lacking an amino acid such as proline in conjunction with E. coli unable to synthesize it were commonly used by geneticists before the emergence of genomics to map bacterial chromosomes.

Selective growth media are also used in cell culture to ensure the survival or proliferation of cells with certain properties, such asantibiotic resistance or the ability to synthesize a certain metabolite. Normally, the presence of a specific gene or an allele of a gene confers upon the cell the ability to grow in the selective medium. In such cases, the gene is termed a marker.

Selective growth media for eukaryotic cells commonly contain neomycin to select cells that have been successfully transfected with a plasmid carrying the neomycin resistance gene as a marker. Gancyclovir is an exception to the rule as it is used to specifically kill cells that carry its respective marker, the Herpes simplex virus thymidine kinase (HSV TK).

Some examples of selective media include:

Eosin methylene blue (EMB) contains dyes that are toxic for Gram positive bacteria and bile salt

which is toxic for Gram negative bacteria other than coliforms. EMB is the selective and

differential medium for coliforms

YM (yeast and mold) which has a low pH, deterring bacterial growth

MacConkey agar for Gram-negative bacteria

Hektoen enteric agar (HE) which is selective for Gram-negative bacteria

Mannitol salt agar (MSA) which is selective for Gram-positive bacteria and differential for

mannitol

Terrific Broth (TB) is used with glycerol in cultivating recombinant strains of Escherichia coli.

Xylose lysine desoxyscholate (XLD), which is selective for Gram-negative bacteria

Buffered charcoal yeast extract agar, which is selective for certain gram-negative bacteria,

especially Legionella pneumophila

Baird–Parker agar for Gram-positive Staphylococci.

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Differential media Differential media or indicator media distinguish one microorganism type from another growing on the same media. [7] This type of media uses the biochemical characteristics of a microorganism growing in the presence of specific nutrients or indicators (such asneutral red, phenol red, eosin y, or methylene blue) added to the medium to visibly indicate the defining characteristics of a microorganism. This type of media is used for the detection of microorganisms and by molecular biologists to detect recombinant strains of bacteria.

Examples of differential media include:

Blood agar (used in strep tests), which contains bovine heart blood that becomes

transparent in the presence of hemolyticStreptococcus

Eosin methylene blue (EMB), which is differential for lactose fermentation

MacConkey (MCK), which is differential for lactose fermentation

Mannitol salt agar (MSA), which is differential for mannitol fermentation

X-gal plates, which are differential for lac operon mutants.

Transport media Transport media should fulfill the following criteria:

temporary storage of specimens being transported to the laboratory for cultivation.

maintain the viability of all organisms in the specimen without altering their concentration.

contain only buffers and salt.

lack of carbon, nitrogen, and organic growth factors so as to prevent microbial

multiplication.

transport media used in the isolation of anaerobes must be free of molecular oxygen.

Examples of transport media include:

Thioglycolate broth for strict anaerobes.

Stuart transport medium - a non-nutrient soft agar gel containing a reducing agent to

prevent oxidation, and charcoal to neutralise

Certain bacterial inhibitors- for gonococci, and buffered glycerol saline for enteric bacilli.

Krishna Rama(KR) medium for V. cholerae.

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Basic Constituents of Media

Inorganic salts

Carbohydrates

Amino Acids

Vitamins

Fatty acids and lipids

Proteins and peptides

Serum

Trace Elements.

Each type of constituent performs a specifi c function as outlined below:

Inorganic Salts

The inclusion of inorganic salts in media performs several functions. Primarily they help to retain the osmotic balance of the cells and help regulate membrane potential by provision of sodium, potassium and calcium ions. All of these are required in the cell matrix for cell attachment and as enzyme cofactors.

Buffering Systems

Most cells require pH conditions in the range 7.2-7.4 and close control of pH is essential for optimum culture conditions. There are major variations to this optimum. Fibroblasts prefer a higher pH (7.4-7.7) whereas, continuous transformed cell lines require more acid conditions pH (7.0-7.4).

Regulation of pH is particularly important immediately following cell seeding when a new culture is establishing and is usually achieved by one of two buffering systems; (i) a “natural” buffering system where gaseous CO2 balances with the CO3/HCO3 content of the culture medium and (ii) chemical buffering using a zwitterion called HEPES.

Cultures using natural bicarbonate /CO2 buffering systems need to be maintained in an atmosphere of 5-10% CO2 in air usually supplied in a CO2 incubator. Bicarbonate/CO2 is low cost, non-toxic and also provides other chemical benefi ts to the cells.

HEPES has superior buffering capacity in the pH range 7.2-7.4 but is relatively expensive and can be toxic to some cell types at higher concentrations. HEPES buffered cultures do not require a controlled gaseous atmosphere.

Most commercial culture media include phenol red as a pH indicator so that the pH status of the medium is constantly indicated by the colour. Usually the culture medium should be changed/replenished if the colour turns yellow (acid) or purple (alkali).

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Carbohydrates

The main source of energy is derived from carbohydrates generally in the form of sugars. The major sugars used are glucose and galactose, however, some media contain maltose or fructose. The concentration of sugar varies from basal media containing 1g/L to 4.5g/L in some more complex media. Media containing the higher concentration of sugars are able to support the growth of a wider range of cell types.

Amino Acids

Amino acids are the building blocks of proteins. ‘Essential’ amino acids must be added to culture media as cells are not able to synthesize these themselves. The concentration of amino acids in the culture medium will determine the maximum cell density that can be achieved - once depleted the cells will no longer be able to proliferate.

In relation to cell culture, glutamine, an essential amino acid, is particularly signifi cant. In liquid media or stock solutions glutamine degrades relatively rapidly. Optimal cell performance usually requires supplementation of the media with glutamine prior to use.

Adding supplements of non-essential amino acids to media both stimulates growth and prolongs the viability of the cells in culture.

Vitamins

Serum is an important source of vitamins in cell culture. However, many media are also enriched with vitamins making them consistently more suitable for a wider range of cell lines. Vitamins are precursors for numerous co-factors. Many vitamins, especially B group vitamins, are necessary for cell growth and proliferation and for some lines the presence of B12 is essential. Some media also have increased levels of vitamins A and E. The vitamins commonly used in media include ribofl avin, thiamine and biotin.

Proteins and Peptides

These are particularly important in serum free media. The most common proteins and peptides include albumin, transferrin, fi bronectin and fetuin and are used to replace those normally present through the addition of serum to the medium.

Fatty Acids and Lipids

Like proteins and peptides these are important in serum free media since they are normally present in serum e.g. cholesterol and steroids essential for specialised cells.

Trace Elements

These include trace elements such as zinc, copper, selenium and tricarboxylic acid intermediates. Selenium is a detoxifi er and helps remove oxygen free radicals.

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Preparation of Media

Whilst all media may be made from the basic ingredients this is time consuming and may predispose to contamination. For convenience most media are available as ready mixed powders or as 10x and 1x liquid media. All commonly used media are listed in the Sigma-Aldrich Life Science Catalogue. If powder or 10x media are purchased it is essential that the water used to reconstitute the powder or dilute the concentrated liquid is free from mineral, organic and microbial contaminants. It must also be pyrogen free and of tissue culture grade. In most cases water prepared by reverse osmosis and resin cartridge purifi cation with a fi nal resistance of 16-18MΩ is suitable. Once prepared, the media should be fi lter sterilised before use.

Serum

Serum is a complex mix of albumins, growth factors and growth inhibitors and is probably one of the most important components of cell culture medium. The most commonly used serum is foetal bovine serum (FBS). Other types of serum are available including newborn calf serum and horse serum. The quality, type and concentration of serum can all affect the growth of cells and it is therefore important to screen batches of serum for their ability to support the growth of cells. In addition, there are other tests that may be used to aid the selection of a batch of serum including cloning effi ciency, plating effi ciency and the preservation of cell characteristics.

Serum is also able to increase the buffering capacity of cultures that can be important for slow growing cells or where the seeding density is low (e.g. cell cloning experiments). It also helps to protect against mechanical damage which may occur in stirred cultures or whilst using a cell scraper.

A further advantage of serum is the wide range of cell types with which it can be used despite the varying requirements of different cultures in terms of growth factors. In addition, serum is able to bind and neutralise toxins. However, serum is subject to batch-to-batch variation that makes standardisation of production protocols difficult.

There is also a risk of contamination associated with the use of serum. These risks can be minimised by obtaining serum from a reputable source since suppliers of large quantities of serum perform a battery of quality control tests and supply a certifi cate of analysis with the serum. In particular, serum is screened for the presence of bovine viral diarrhoea virus (BVDV) and mycoplasma. Heat inactivation of serum (incubation at 56°C for 30 minutes) can help to reduce the risk of contamination since some viruses are inactivated by this process. However, the routine use of heat inactivated serum is not an absolute requirement for cell culture. The use of serum also has a cost implication not only in terms of medium formulation but also in downstream processing. A 10% FBS supplement contributes 4.8mg of protein per millilitre of culture fl uid which complicates downstream processing procedures such as protein purification.

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Guidelines for Serum Use- Foetal bovine serum (FBS) has been used to prepare a number of biologicals and has an excellent record of safety. The recognition of Bovine Spongiform Encephalopathy (BSE) in 1986 and its subsequent spread into continental Europe along side the announcement of the probable link between BSE and a new variant of Creutzfeldt Jacob disease in Humans stimulated an increased concern about safe sourcing of all bovine materials. In 1993, the Food and Drug Administration (FDA) "recommended against the use of bovine derived materials from cattle which have resided in, or originated from countries where BSE has been diagnosed”.

The current European Union (EU) guidelines on viral safety focus on sourcing, testing and paying particular attention to the potential risk of cross contamination during slaughtering or collection of the starting tissue.

As far as BSE is concerned, the EU guidance on minimising the risk of BSE transmission via medicinal products, EMEA/410/01 Rev. 2, recommends the main measures to be implemented in order to establish the safety of bovine material. Similarly the focus is on geographical origin, the age of the animals, the breeding and slaughtering conditions, the tissue to be used and the conditions of its processing.

The use of FBS in production processes of medicinal products is acceptable provided good documentation on sourcing, age of the animals and testing for the absence of adventitious agents is submitted. All responsible suppliers of FBS for bio-pharmaceutical applications will provide such documentation.

Regulatory requirements in Europe stress the importance of justifying the use of material of bovine, caprine or ovine origin in the production of pharmaceutical products. Thus, although FBS has been used for many years in the production process of many medicinal products such as viral vaccines and recombinant DNA products, at present there is a justifi ed trend to remove all material of animal origin from manufacturing processes. Sigma- Aldrich has recognised this growing trend and works closely with customers to optimise animal free media formulations to meet each customer’s cell culture requirements. Serum-free cell lines that have been adapted to media that do not contain serum are available from ECACC.

The United States Department of Agriculture (USDA) regulates all products that contain a primary component of animal origin. With specifi c reference to serum the USDA has declared that for materials which fall under their jurisdiction, only biological products manufactured using serum from approved countries of origin will be allowed in to USA.

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Origin of Serum

ECACC only uses serum of Zone 1 origin, sterile fi ltered and cell culture tested. Zone 1 countries have BSE –free status such as the USA, Canada, Australia and New Zealand. It is essential to check the source country of the serum used and their Zone status. Sera from Mexico and Central American countries may require additional documentation to prove the geographical region of the donor herd to ensure BSE-free status. This is very important if the intended use of the serum is in the production of medicinal or other products being sent to the USA.

Serum from a reputable supplier should have undergone various quality control tests which will be listed in the product information sheet. Most serum products are cell culture tested including growth promotion, cloning effi ciency and plating effi ciency tests.

Standard tests performed on serum commonly include tests to determine the presence and/or level of the following:

Sterility

Virus Contamination

Mycoplasma Contamination

Endotoxin

Haemoglobin

Total Protein

Immunoglobulin

Hormone Testing

pH (at room temperature)

Osmolality

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Laminar flow

A sphere in Stokes flow, at very low Reynolds number. An object moving through a fluid experiences a force in the

direction opposite to its motion.

In fluid dynamics, laminar flow (or streamline flow) occurs when a fluid flows in parallel layers, with no disruption between the layers.[1 ] At low velocities, the fluid tends to flow without lateral mixing, and adjacent layers slide past one another like playing cards. There are no cross-currents perpendicular to the direction of flow, nor eddies or swirls of fluids.[2] In laminar flow, the motion of the particles of the fluid is very orderly with all particles moving in straight lines parallel to the pipe walls.[3] Laminar flow is a flow regime characterized by high momentum diffusion and low momentum convection.

When a fluid is flowing through a closed channel such as a pipe or between two flat plates, either of two types of flow may occur depending on the velocity of the fluid: laminar flow or turbulent flow. Laminar flow tends to occur at lower velocities, below a threshold at which it becomes turbulent. Turbulent flow is a less orderly flow regime that is characterised by eddies or small packets of fluid particles which result in lateral mixing.[2 ] In non-scientific terms, laminar flow is smooth while turbulent flow is rough.

Relationship with the Reynolds number The type of flow occurring in a fluid in a channel is important in fluid dynamics problems. The dimensionless Reynolds number is an important parameter in the equations that describe whether flow conditions lead to laminar or turbulent flow. The Reynolds number delimiting laminar and turbulent flow depends on the particular flow geometry, and moreover, the transition from laminar to turbulent flow can be sensitive to disturbance levels and imperfections present in a given configuration.

In the case of flow through a straight pipe with a circular cross-section, at a Reynolds number below a critical value of approximately 2040,[4] fluid motion will ultimately be laminar, whereas at larger Reynolds numbers, the flow can be turbulent. When the Reynolds number is much less than

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1, Stokes flow occurs. This is an extreme case of laminar flow whereby viscous (frictional) effects are much greater than inertial forces.

A common application of laminar flow is in the smooth flow of a viscous liquid through a tube or pipe. In that case, the velocity of flow varies from zero at the walls to a maximum along the cross-sectional centre of the vessel. The flow profile of laminar flow in a tube can be calculated by dividing the flow into thin cylindrical elements and applying the viscous force to them.[5]

Another example is the flow of air over an aircraft wing. The boundary layer is a very thin sheet of air lying over the surface of the wing (and all other surfaces of the aircraft). Because air has viscosity, this layer of air tends to adhere to the wing. As the wing moves forward through the air, the boundary layer at first flows smoothly over the streamlined shape of the airfoil. Here, the flow is laminar and the boundary layer is a laminar layer. Prandtl applied the concept of the laminar boundary layer to airfoils in 1904. [6][7 ]

Laminar flow barriers

Experimental chamber for studyingchemotaxis in response to laminar flow.

Laminar airflow is used to separate volumes of air, or prevent airborne contaminants from entering an area. Laminar flow hoods are used to exclude contaminants from sensitive processes in science, electronics and medicine. Air curtains are frequently used in commercial settings to keep heated or refrigerated air from passing through doorways. A laminar flow reactor (LFR) is a reactor that uses laminar flow to study chemical reactions and process mechanisms.

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Laminar flow reactor Laminar flow reactor (LFR) is a type of chemical reactor that uses laminar flow to control reaction rate, and/or reaction distribution. LFR is generally a long tube with constant diameter that is kept at constant temperature. Reactants are injected at one end and products are collected and monitored at the other.[1] Laminar flow reactors are often used to study an isolated elementary reaction or multi-step reaction mechanism.

Overview

Laminar Flow Reactor employs the characteristics of laminar flow to achieve various research purposes. For instance, LFRs can be used to study fluid dynamics in chemical reactions, or they can be utilized to generate special chemical structures such as carbon nanotubes. One feature of the LFR is that the residence time (The time interval during which the chemicals stay in the reactor) of the chemicals in the reactor can be varied by either changing the distance between the reactant input point and the point at which the product/sample is taken, or by adjusting the velocity of the gas/fluid. Therefore the benefit of a laminar flow reactor is that the different factors that may affect a reaction can be easily controlled and adjusted throughout an experiment.

Means of Analyzing Reactants in LFR

Means of analyzing the reaction include using a probe that enters into the reactor; or more accurately, sometimes one can utilize non-intrusive optical methods (e.g. usespectrometer to identify and analyze contents) to study reactions in the reactor. Moreover, taking the entire sample of the gas/fluid at the end of the reactor and collecting data may be useful as well.[1] Using methods mentioned above, various data such as concentration, flow velocity etc. can be monitored and analyzed.

Flow Velocity in LFR

Fluids or gases with controlled velocity pass through a laminar flow reactor in a fashion of laminar flow. That is, streams of fluids or gases slide over each other like cards. When analyzing fluids with same the viscosity ("thickness" or "stickiness") but different velocity, fluids is typically characterized into two types of flows: laminar flow and turbulent flow. Compared to turbulent flow, laminar flow tends to have a lower velocity and is generally at a lower Reynolds number. Turbulent flow, on the other hand, is irregular and travels at a higher speed. Therefore the flow velocity of a turbulent flow on one cross section is often assumed to be constant, or "flat". The "non-flat" flow velocity of laminar flow helps explain the mechanism of a LFR. For the fluid/gas moving of a LFR, the velocity near the center of the pipe is higher than the fluids near the wall of the pipe. Thus, the velocity distribution of the reactants tends to decrease from the center to the wall.

Residence Time Distribution(RTD)

The velocity near the center of the pipe is higher than the fluids near the wall of the pipe. Thus, the velocity distribution of the reactants tends to be higher in the center and lower on the side. Consider fluid being pumped through a LFR at constant velocity from the inlet, and the concentration of the fluid is monitored at the outlet. The graph of the residence time distribution should look like a negative slope with positive concavity. And the graph is modeled by the function:

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E(t)=0 if t is smaller than τ/2; E(t)=τ^2/2t^3 if t is greater than or equal to τ/2.[2 ] Notice that the graph has the τ value of zero initially, this is simply because it takes sometime for the substance to travel through the reactor. When the material is starting to reach the outlet, the concentration drastically increases, and it gradually decreases as time proceeds.

Characteristics

The laminar flows inside of a LFR has the unique characteristic of flowing in a parallel fashion without disturbing one another. The velocity of the fluid or gas will naturally decrease as it gets closer to the wall and farther from the center. Therefore the reactants have a decreasing residence time in the LFR from the center to the side. A gradually decreasing residence time gives researchers a clear layout of the reaction at different times. Besides, when studying reactions in LFR, radial gradients in velocity, composition and temperature are significant.[3] In other words, in other reactors where laminar flow is not significant, for instance, in a plug flow reactor, velocity of the object is assumed to be the same on one cross section since the flows are mostly turbulent. In a laminar flow reactor, velocity is significantly different at various points on the same cross section. Therefore the velocity differences throughout the reactor need to be taken into consideration when working with a LFR.

Research

Various researches pertaining to the modeling of LFR and formations of substances within a LFR have been done over the past decades. For instance, the formation of Single-walled carbon nanotube was investigated in a LFR.[4] As another example, conversion from methane to higher hydrocarbons have been studied in a laminar flow reactor.[5]

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Incubator (culture)

A Bacteriological incubator

Interior of aCO2 incubator used in cell culture

In biology, an incubator is a device used to grow and maintain microbiological cultures or cell cultures. The incubator maintains optimal temperature,humidity and other conditions such as the carbon dioxide (CO2) and oxygen content of the atmosphere inside. Incubators are essential for a lot of experimental work in cell biology, microbiology and molecular biology and are used to culture both bacterial as well as eukaryotic cells.

Incubators are also used in the poultry industry to act as a substitute for hens. This often results in higher hatch rates due to the ability to control both temperature and humidity. Various brands of incubators are commercially available to breeders.

The simplest incubators are insulated boxes with an adjustable heater, typically going up to 60 to 65 °C (140 to 150 °F), though some can go slightly higher (generally to no more than 100 °C). The most commonly used temperature both for bacteria such as the frequently used E. coli as well as for mammalian cells is approximately 37 °C, as these organisms grow well under such conditions. For other organisms used in biological experiments, such as the budding yeast Saccharomyces cerevisiae, a growth temperature of 30 °C is optimal.

More elaborate incubators can also include the ability to lower the temperature (via refrigeration), or the ability to control humidity or CO2 levels. This is important in the cultivation of mammalian cells, where the relative humidity is typically >80% to prevent evaporation and a slightly acidic pH is achieved by maintaining a CO2 level of 5%.

Louis Pasteur used the small opening underneath his staircase as an incubator.

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Fluorescence microscope

An upright fluorescence microscope (Olympus BX61) with the fluorescent filter cube turret above the objective

lenses, coupled with a digital camera.

A fluorescence microscope is an optical microscope that uses fluorescence and phosphorescence instead of, or in addition to, reflectionand absorption to study properties of organic or inorganic substances.[1][2] The "fluorescence microscope" refers to any microscope that uses fluorescence to generate an image, whether it is a more simple set up like an epifluorescence microscope, or a more complicated design such as a confocal microscope, which uses optical sectioning to get better resolution of the fluorescent image.

On October 8, 2014, the Nobel Prize in Chemistry was awarded to Eric Betzig, William Moerner and Stefan Hell for "the development of super-resolved fluorescence microscopy," which brings "optical microscopy into the nanodimension".[3][4]

Principle The specimen is illuminated with light of a specific wavelength (or wavelengths) which is absorbed by the fluorophores, causing them to emit light of longer wavelengths (i.e., of a different color than the absorbed light). The illumination light is separated from the much weaker emitted fluorescence through the use of a spectral emission filter. Typical components of a fluorescence microscope are a light source (xenon arc lamp or mercury-vapor lamp are common; more advanced forms are high-power LEDs and lasers), theexcitation filter, the dichroic mirror (or dichroic beamsplitter), and the emission filter (see figure below). The filters and the dichroic are chosen to match the spectral excitation and emission characteristics of the fluorophore used to label the specimen.[1] In this manner, the distribution of a single fluorophore (color) is imaged at a time. Multi-color images of several types of fluorophores must be composed by combining several single-color images.[1]

Most fluorescence microscopes in use are epifluorescence microscopes, where excitation of the fluorophore and detection of the fluorescence are done through the same light path (i.e. through the

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objective). These microscopes are widely used in biology and are the basis for more advanced microscope designs, such as the confocal microscope and the total internal reflection fluorescence microscope (TIRF).

Epifluorescence microscopy

Schematic of a fluorescence microscope.

The majority of fluorescence microscopes, especially those used in the life sciences, are of the epifluorescence design shown in the diagram. Light of the excitation wavelength is focused on the specimen through the objective lens. The fluorescence emitted by the specimen is focused to the detector by the same objective that is used for the excitation which for greatest sensitivity will have a very highnumerical aperture. Since most of the excitation light is transmitted through the specimen, only reflected excitatory light reaches the objective together with the emitted light and the epifluorescence method therefore gives a high signal-to-noise ratio. An additional barrier filter between the objective and the detector can filter out the remaining excitation light from fluorescent light.

Light sources

Fluorescence microscopy requires intense, near-monochromatic, illumination which some widespread light sources, like halogen lampscannot provide. Four main types of light source are used, including xenon arc lamps or mercury-vapor lamps with an excitation filter,lasers, supercontinuum sources, and high-power LEDs. Lasers are most widely used for more complex fluorescence microscopy techniques like confocal microscopy and total internal reflection fluorescence microscopy while xenon lamps, and mercury lamps, and LEDs with a dichroic excitation filter are commonly used for widefield epifluorescence microscopes.

Sample preparation

In order for a sample to be suitable for fluorescence microscopy it must be fluorescent. There are several methods of creating a fluorescent sample; the main techniques are labelling with fluorescent stains or, in the case of biological samples, expression of a fluorescent protein. Alternatively the intrinsic fluorescence of a sample (i.e., autofluorescence) can be used.[1 ] In the life

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sciences fluorescence microscopy is a powerful tool which allows the specific and sensitive staining of a specimen in order to detect the distribution of proteins or other molecules of interest. As a result there is a diverse range of techniques for fluorescent staining of biological samples.

Biological fluorescent stains

Many fluorescent stains have been designed for a range of biological molecules. Some of these are small molecules which are intrinsically fluorescent and bind a biological molecule of interest. Major examples of these are nucleic acid stains like DAPI and Hoechst (excited by UV wavelength light) and DRAQ5 and DRAQ7 (optimally excited by red light) which all bind the minor groove of DNA, thus labelling the nuclei of cells. Others are drugs or toxins which bind specific cellular structures and have been derivatised with a fluorescent reporter. A major example of this class of fluorescent stain is phalloidin which is used to stain actin fibres in mammalian cells.

There are many fluorescent molecules called fluorophores or fluorochromes such as fluorescein, Alexa Fluors or DyLight 488, which can be chemically linked to a different molecule which binds the target of interest within the sample.

Immunofluorescence Immunofluorescence is a technique which uses the highly specific binding of an antibody to its antigen in order to label specific proteins or other molecules within the cell. A sample is treated with a primary antibody specific for the molecule of interest. A fluorophore can be directly conjugated to the primary antibody. Alternatively a secondary antibody, conjugated to a fluorophore, which binds specifically to the first antibody can be used. For example a primary antibody raised in a mouse which recognises tubulin combined with a secondary anti-mouse antibody derivatised with a fluorophore could be used to label microtubules in a cell.

Fluorescent proteins The modern understanding of genetics and the techniques available for modifying DNA allow scientists to genetically modify proteins to also carry a fluorescent protein reporter. In biological samples this allows a scientist to directly make a protein of interest fluorescent. The protein location can then be directly tracked, including in live cells.

Limitations

Fluorophores lose their ability to fluoresce as they are illuminated in a process called photobleaching. Photobleaching occurs as the fluorescent molecules accumulate chemical damage from the electrons excited during fluorescence. Photobleaching can severely limit the time over which a sample can be observed by fluorescent microscopy. Several techniques exist to reduce photobleaching such as the use of more robust fluorophores, by minimizing illumination, or by using photoprotective scavenger chemicals.

Fluorescence microscopy with fluorescent reporter proteins has enabled analysis of live cells by fluorescence microscopy, however cells are susceptible to phototoxicity, particularly with short wavelength light. Furthermore fluorescent molecules have a tendency to generate reactive chemical species when under illumination which enhances the phototoxic effect.

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Unlike transmitted and reflected light microscopy techniques fluorescence microscopy only allows observation of the specific structures which have been labeled for fluorescence. For example, observing a tissue sample prepared with a fluorescent DNA stain by fluorescent microscopy only reveals the organisation of the DNA within the cells and reveals nothing else about the cell morphologies.

Sub-diffraction techniques

The wave nature of light limits the size of the spot to which light can be focused due to the diffraction limit. This limitation was described in the 19th century by Ernst Abbe and limits an optical microscope's resolution to approximately half of the wavelength of the light used. Fluorescence microscopy is central to many techniques which aim to reach past this limit by specialised optical configurations.

Several improvements in microscopy techniques have been invented in the 20th century and have resulted in increased resolution and contrast to some extent. However they did not overcome the diffraction limit. In 1978 first theoretical ideas have been developed to break this barrier by using a 4Pi microscope as a confocal laser scanning fluorescence microscope where the light is focused ideally from all sides to a common focus which is used to scan the object by 'point-by-point' excitation combined with 'point-by-point' detection.[5] However, the first experimental demonstration of the 4pi microscope took place in 1994.[6] 4Pi microscopy maximizes the amount of available focusing directions by using two opposing objective lenses or Multi-photon microscopy using redshifted light and multi-photon excitation.

The first technique to really achieve a sub-diffraction resolution was STED microscopy, proposed in 1994. This method and all techniques following the RESOLFT concept rely on a strong non-linear interaction between light and fluorescing molecules. The molecules are driven strongly between distinguishable molecular states at each specific location, so that finally light can be emitted at only a small fraction of space, hence an increased resolution.

As well in the 1990s another super resolution microscopy method based on wide field microscopy has been developed. Substantially improved size resolution of cellularnanostructures stained with a fluorescent marker was achieved by development of SPDM localization microscopy and the structured laser illumination (spatially modulated illumination, SMI).[7] Combining the principle of SPDM with SMI resulted in the development of the Vertico SMI microscope.[8][9 ] Single molecule detection of normal blinkingfluorescent dyes like Green fluorescent protein (GFP) can be achieved by using a further development of SPDM the so-called SPDMphymod technology which makes it possible to detect and count two different fluorescent molecule types at the molecular level (this technology is referred to as two-color localization microscopy or 2CLM).[10]

Alternatively, the advent of photoactivated localization microscopy could achieve similar results by relying on blinking or switching of single molecules, where the fraction of fluorescing molecules is very small at each time. This stochastic response of molecules on the applied light corresponds also to a highly nonlinear interaction, leading to subdiffraction resolution.

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ELISA enzyme-linked immunosorbent assay (ELISA) The enzyme-linked immunosorbent assay (ELISA) (/ɨˈlaɪzə/, /ˌiːˈlaɪzə/) is a test that uses antibodies and color change to identify a substance.

ELISA is a popular format of "wet-lab" type analytic biochemistry assay that uses a solid-phase enzyme immunoassay (EIA) to detect the presence of a substance, usually an antigen, in a liquid sample or wet sample.

The ELISA has been used as a diagnostic tool in medicine and plant pathology, as well as a quality-control check in various industries.

Antigens from the sample are attached to a surface. Then, a further specific antibody is applied over the surface so it can bind to the antigen. This antibody is linked to an enzyme, and, in the final step, a substance containing the enzyme's substrate is added. The subsequent reaction produces a detectable signal, most commonly a color change in the substrate.

Performing an ELISA involves at least one antibody with specificity for a particular antigen. The sample with an unknown amount of antigen is immobilized on a solid support (usually a polystyrene microtiter plate) either non-specifically (via adsorption to the surface) or specifically (via capture by another antibody specific to the same antigen, in a "sandwich" ELISA). After the antigen is immobilized, the detection antibody is added, forming a complex with the antigen. The detection antibody can be covalently linked to an enzyme, or can itself be detected by a secondary antibody that is linked to an enzyme through bioconjugation. Between each step, the plate is typically washed with a mild detergent solution to remove any proteins or antibodies that are non-specifically bound. After the final wash step, the plate is developed by adding an enzymatic substrate to produce a visible signal, which indicates the quantity of antigen in the sample.

Of note, ELISA can perform other forms of ligand binding assays instead of strictly "immuno" assays, though the name carried the original "immuno" because of the common use and history of development of this method. The technique essentially requires any ligating reagent that can be immobilized on the solid phase along with a detection reagent that will bind specifically and use an enzyme to generate a signal that can be properly quantified. In between the washes, only the ligand and its specific binding counterparts remain specifically bound or "immunosorbed" by antigen-antibody interactions to the solid phase, while the nonspecific or unbound components are washed away. Unlike other spectrophotometric wet lab assay formats where the same reaction well (e.g. a cuvette) can be reused after washing, the ELISA plates have the reaction products immunosorbed on the solid phase which is part of the plate, and so are not easily reusable.

Principle As an analytic biochemistry assay, ELISA involves detection of an "analyte" (i.e. the specific substance whose presence is being quantitatively or qualitatively analyzed) in a liquid sample by a method that continues to use liquid reagents during the "analysis" (i.e. controlled sequence of biochemical reactions that will generate a signal which can be easily quantified and interpreted as a measure of the amount of analyte in the sample) that stays liquid and remains inside a reaction chamber or well needed to keep the reactants contained; It is opposed to "dry lab" that can use dry strips – and even if the sample is liquid (e.g. a measured small drop), the final detection step in

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"dry" analysis involves reading of a dried strip by methods such as reflectometry and does not need a reaction containment chamber to prevent spillover or mixing between samples.

As a heterogenous assay, ELISA separates some component of the analytical reaction mixture by adsorbing certain components onto a solid phase which is physically immobilized. In ELISA, a liquid sample is added onto a stationary solid phase with special binding properties and is followed by multiple liquid reagents that are sequentially added, incubated and washed followed by some optical change (e.g. color development by the product of an enzymatic reaction) in the final liquid in the well from which the quantity of the analyte is measured. The qualitative "reading" usually based on detection of intensity of transmitted light by spectrophotometry, which involves quantitation of transmission of some specific wavelength of light through the liquid (as well as the transparent bottom of the well in the multiple-well plate format). The sensitivity of detection depends on amplification of the signal during the analytic reactions. Since enzyme reactions are very well known amplification processes, the signal is generated by enzymes which are linked to the detection reagents in fixed proportions to allow accurate quantification – thus the name "enzyme linked".

The analyte is also called the ligand because it will specifically bind or ligate to a detection reagent, thus ELISA falls under the bigger category of ligand binding assays. The ligand-specific binding reagent is "immobilized", i.e., usually coated and dried onto the transparent bottom and sometimes also side wall of a well (the stationary "solid phase'/"solid substrate" here as opposed to solid microparticle/beads that can be washed away), which is usually constructed as a multiple-well plate known as the "ELISA plate". Conventionally, like other forms of immunoassays, the specificity of antigen-antibody type reaction is used because it is easy to raise an antibody specifically against an antigen in bulk as a reagent. Alternatively, if the analyte itself is an antibody, its target antigen can be used as the binding reagent.

History Before the development of the ELISA, the only option for conducting an immunoassay was radioimmunoassay, a technique using radioactively labeled antigens or antibodies. In radioimmunoassay, the radioactivity provides the signal, which indicates whether a specific antigen or antibody is present in the sample. Radioimmunoassay was first described in a scientific paper by Rosalyn Sussman Yalow and Solomon Berson published in 1960.[1 ]

Because radioactivity poses a potential health threat, a safer alternative was sought. A suitable alternative to radioimmunoassay would substitute a nonradioactive signal in place of the radioactive signal. When enzymes (such as horseradish peroxidase) react with appropriate substrates (such as ABTS or TMB), a change in color occurs, which is used as a signal. However, the signal has to be associated with the presence of antibody or antigen, which is why the enzyme has to be linked to an appropriate antibody. This linking process was independently developed by Stratis Avrameas and G. B. Pierce.[2] Since it is necessary to remove any unbound antibody or antigen by washing, the antibody or antigen has to be fixed to the surface of the container; i.e., the immunosorbent must be prepared. A technique to accomplish this was published by Wide and Jerker Porath in 1966. [3]

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Paramedic assistant prepares the analyses in ELISA laboratory

In 1971, Peter Perlmann and Eva Engvall at Stockholm University in Sweden, and Anton Schuurs and Bauke van Weemen in the Netherlands independently published papers that synthesized this knowledge into methods to perform EIA/ELISA.[4][5 ]

Traditional ELISA typically involves chromogenic reporters and substrates that produce some kind of observable color change to indicate the presence of antigen or analyte. Newer ELISA-like techniques use fluorogenic, electro-chemiluminescent, and quantitative PCRreporters to create quantifiable signals. These new reporters can have various advantages, including higher sensitivities andmultiplexing.[6 ][7] In technical terms, newer assays of this type are not strictly ELISAs, as they are not "enzyme-linked", but are instead linked to some nonenzymatic reporter. However, given that the general principles in these assays are largely similar, they are often grouped in the same category as ELISAs.

In 2012 an ultrasensitive, enzyme-based ELISA test using nanoparticles as a chromogenic reporter was able to give a naked-eye colour signal from the detection of mere attograms of analyte. A blue color appears for positive results and red color for negative. Note that this detection only can confirm the presence or the absence of analyte not the actual concentration.[8]

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Types of ELISA Direct ELISA

Direct ELISA diagram

The steps of direct ELISA follows the mechanism below:-

A buffered solution of the antigen to be tested for is added to each well of a microtiter plate,

where it is given time to adhere to the plastic through charge interactions.

A solution of nonreacting protein, such as bovine serum albumin or casein, is added to well

(usually 96-well plates) in order to cover any plastic surface in the well which remains

uncoated by the antigen.

The primary antibody with an attached (conjugated) enzyme is added, which binds

specifically to the test antigen coating the well.

A substrate for this enzyme is then added. Often, this substrate changes color upon reaction

with the enzyme.

The higher the concentration of the primary antibody present in the serum, the stronger the

color change. Often, a spectrometer is used to give quantitative values for color strength.

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The enzyme acts as an amplifier; even if only few enzyme-linked antibodies remain bound, the enzyme molecules will produce many signal molecules. Within common-sense limitations, the enzyme can go on producing color indefinitely, but the more antibody is bound, the faster the color will develop. A major disadvantage of the direct ELISA is the method of antigen immobilization is not specific; when serum is used as the source of test antigen, all proteins in the sample may stick to the microtiter plate well, so small concentrations of analyte in serum must compete with other serum proteins when binding to the well surface. The sandwich or indirect ELISA provides a solution to this problem, by using a "capture" antibody specific for the test antigen to pull it out of the serum's molecular mixture.

ELISA may be run in a qualitative or quantitative format. Qualitative results provide a simple positive or negative result (yes or no) for a sample. The cutoff between positive and negative is determined by the analyst and may be statistical. Two or three times the standard deviation (error inherent in a test) is often used to distinguish positive from negative samples. In quantitative ELISA, the optical density (OD) of the sample is compared to a standard curve, which is typically a serial dilution of a known-concentration solution of the target molecule. For example, if a test sample returns an OD of 1.0, the point on the standard curve that gave OD = 1.0 must be of the same analyte concentration as the sample.

The use and meaning of the names "direct ELISA" and "indirect ELISA" differs in the literature and on web sites depending on the context of the experiment. When the presence of an antigen is analyzed, the name "direct ELISA" refers to an ELISA in which only a labelled primary antibody is used, and the term "indirect ELISA" refers to an ELISA in which the antigen is bound by the primary antibody which then is detected by a labelled secondary antibody. In the latter case a sandwich ELISA is clearly distinct from an indirect ELISA. When the 'primary' antibody is of interest, e.g. in the case of immunization analyses, this antibody is directly detected by the secondary antibody and the term "direct ELISA" applies to a setting with two antibodies.

Sandwich ELISA

A sandwich ELISA. (1) Plate is coated with a capture antibody; (2) sample is added, and any

antigen present binds to capture antibody; (3) detecting antibody is added, and binds to antigen;

(4) enzyme-linked secondary antibody is added, and binds to detecting antibody; (5) substrate is

added, and is converted by enzyme to detectable form.

A "sandwich" ELISA, is used to detect sample antigen. The steps are:

1. A surface is prepared to which a known quantity of capture antibody is bound.

2. Any nonspecific binding sites on the surface are blocked.

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3. The antigen-containing sample is applied to the plate, and captured by antibody.

4. The plate is washed to remove unbound antigen.

5. A specific antibody is added, and binds to antigen (hence the 'sandwich': the Ag is stuck

between two antibodies). This primary antibody could also be in the serum of a donor to be

tested for reactivity towards the antigen.

6. Enzyme-linked secondary antibodies are applied as detection antibodies that also bind

specifically to the antibody's Fc region (nonspecific).

7. The plate is washed to remove the unbound antibody-enzyme conjugates.

8. A chemical is added to be converted by the enzyme into a color or fluorescent or

electrochemical signal.

9. The absorbance or fluorescence or electrochemical signal (e.g., current) of the plate wells is

measured to determine the presence and quantity of antigen.

The image to the right includes the use of a secondary antibody conjugated to an enzyme, though, in the technical sense, this is not necessary if the primary antibody is conjugated to an enzyme (which would be direct ELISA). However, the use of a secondary-antibody conjugate avoids the expensive process of creating enzyme-linked antibodies for every antigen one might want to detect. By using an enzyme-linked antibody that binds the Fc region of other antibodies, this same enzyme-linked antibody can be used in a variety of situations. Without the first layer of "capture" antibody, any proteins in the sample (including serum proteins) may competitively adsorb to the plate surface, lowering the quantity of antigen immobilized. Use of the purified specific antibody to attach the antigen to the plastic eliminates a need to purify the antigen from complicated mixtures before the measurement, simplifying the assay, and increasing the specificity and the sensitivity of the assay.

A descriptive animation of the application of sandwich ELISA to home pregnancy testing can be found here.

Competitive ELISA A third use of ELISA is through competitive binding. The steps for this ELISA are somewhat different from the first two examples:

1. Unlabeled antibody is incubated in the presence of its antigen (sample).

2. These bound antibody/antigen complexes are then added to an antigen-coated well.

3. The plate is washed, so unbound antibodies are removed. (The more antigen in the sample,

the more Ag-Ab complexes are formed and so there are less unbound antibodies available

to bind to the antigen in the well, hence "competition".)

4. The secondary antibody, specific to the primary antibody, is added. This second antibody is

coupled to the enzyme.

5. A substrate is added, and remaining enzymes elicit a chromogenic or fluorescent signal.

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6. The reaction is stopped to prevent eventual saturation of the signal.

Some competitive ELISA kits include enzyme-linked antigen rather than enzyme-linked antibody. The labeled antigen competes for primary antibody binding sites with the sample antigen (unlabeled). The less antigen in the sample, the more labeled antigen is retained in the well and the stronger the signal.

Commonly, the antigen is not first positioned in the well.

For the detection of HIV antibodies, the wells of microtiter plate are coated with the HIV antigen. Two specific antibodies are used, one conjugated with enzyme and the other present in serum (if serum is positive for the antibody). Cumulative competition occurs between the two antibodies for the same antigen, causing a stronger signal to be seen. Sera to be tested are added to these wells and incubated at 37 °C, and then washed. If antibodies are present, the antigen-antibody reaction occurs. No antigen is left for the enzyme-labelled specific HIV antibodies. These antibodies remain free upon addition and are washed off during washing. Substrate is added, but there is no enzyme to act on it, so positive result shows no color change.

Animated video overview of competitive ELISA.

Multiple and ready to use ELISA A new technique (EP 1 499 894 B1 in EPO Bulletin 25.02.209 N. 2009/09; USPTO 7510687 in USPTO Bulletin 31.03.2009; ZL 03810029.0 in SIPO PRC Bulletin 08.04.2009) uses a solid phase made up of an immunosorbent polystyrene rod with eight to 12 protruding ogives. The entire device is immersed in a test tube containing the collected sample and the following steps (washing, incubation in conjugate and incubation in chromogens) are carried out by dipping the ogives in microwells of standard microplates filled with reagents.

The advantages of this technique are:

1. The ogives can each be sensitized to a different reagent, allowing the simultaneous

detection of different antibodies and/or different antigens for multiple-target assays.

2. The sample volume can be increased to improve the test sensitivity in clinical (blood, saliva,

urine), food (bulk milk, pooled eggs) and environmental (water) samples.

3. One ogive is left unsensitized to measure the nonspecific reactions of the sample.

4. The use of laboratory supplies for dispensing sample aliquots, washing solution and

reagents in microwells is not required, facilitating the development of ready-to-use lab kits

and on-site testing.

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Applications

Human anti-IgG, double antibody sandwich ELISA

Because the ELISA can be performed to evaluate either the presence of antigen or the presence of antibody in a sample, it is a useful tool for determining serum antibody concentrations (such as with the HIV test[9] or West Nile virus). It has also found applications in the foodindustry in detecting potential food allergens, such as milk, peanuts, walnuts, almonds, and eggs[10] and as serological blood test forcoeliac disease.[11 ][12 ] ELISA can also be used in toxicology as a rapid presumptive screen for certain classes of drugs.

Enzyme-linked immunosorbent assay plate

The ELISA was the first screening test widely used for HIV because of its high sensitivity. In an ELISA, a person's serum is diluted 400 times and applied to a plate to which HIV antigens are attached. If antibodies to HIV are present in the serum, they may bind to these HIV antigens. The plate is then washed to remove all other components of the serum. A specially prepared "secondary antibody" — an antibody that binds to other antibodies — is then applied to the plate, followed by another wash. This secondary antibody is chemically linked in advance to an enzyme.

Thus, the plate will contain enzyme in proportion to the amount of secondary antibody bound to the plate. A substrate for the enzyme is applied, and catalysis by the enzyme leads to a change in color or fluorescence. ELISA results are reported as a number; the most controversial aspect of this test is determining the "cut-off" point between a positive and a negative result.

A cut-off point may be determined by comparing it with a known standard. If an ELISA test is used for drug screening at workplace, a cut-off concentration, 50 ng/ml, for example, is established, and a sample containing the standard concentration of analyte will be prepared. Unknowns that generate a stronger signal than the known sample are "positive." Those that generate weaker signal are "negative".

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Dr Dennis E Bidwell and Alister Voller created the ELISA test to detect various kind of diseases, such as malaria, Chagas disease, andJohne's disease.[13] ELISA tests also are used as in in vitro diagnostics in medical laboratories. The other uses of ELISA include:

detection of Mycobacterium antibodies in tuberculosis

detection of rotavirus in feces

detection of hepatitis B markers in serum

detection of enterotoxin of E. coli in feces

detection of HIV antibodies in blood samples

What types of ELISA assays may you use? Four kinds of ELISA here are here illustrated as you may concern:

Direct ELISA

Direct ELISAs involve attachment of the antigen to the solid phase, followed by an enzyme-

labeled antibody. This type of assay generally makes measurement of crude samples

difficult, since contaminating proteins compete for plastic binding sites.

Indirect ELISA

Indirect ELISAs also involve attachment of the antigen to a solid phase, but in this case, the

primary antibody is not labeled. An enzyme-conjugated secondary antibody, directed at the

first antibody, is then added. This format is used most often to detect specific antibodies in

sera.

Competitive ELISA

The third type of ELISA is the Competition Assay, which involves the simultaneous addition

of 'competing' antibodies or proteins. The decrease in signal of samples where the second

antibody or protein is added gives a highly specific result.

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Sandwich ELISA

The last type of assay is the sandwich ELISA. Sandwich ELISAs involve attachment of a

capture antibody to a solid phase support. Samples containing known or unknown antigen

are then added in a matrix or buffer that will minimize attachment to the solid phase. An

enzyme-labeled antibody is then added for detection.

The ELISA method is a benchmark for quantitation of pathological antigens and there are

indeed many variations to this method. ELISAs are adaptable to high-throughput screening

because results are rapid, consistent and relatively easy to analyze. The best results have

been obtained with the sandwich format, utilizing highly purified, prematched capture and

detector antibodies. The resulting signal provides data which is very sensitive and highly

specific.

All ELISAs rely on the specific interaction between an epitope, a small linear or three dimensional sequence of amino acids found on an antigen, and a matching antibody binding site. The antibodies used in an ELISA can be either monoclonal (derived from unique antibody producing cells called hybridomas and capable of specific binding to a single unique epitope) or polyclonal (a pool of antibodies purified from animal sera that are capable of binding to multiple epitopes). There are four basic ELISA formats, allowing for a certain amount of flexibility which can be adjusted based on the antibodies available, the results required, or the complexity of the samples. It is possible to use both monoclonals and polyclonals in an ELISA; however, polyclonals are more typically used for the secondary detection layer in indirect ELISAs, while monoclonal antibodies are more typically used for capture or primary detection of the antigen.

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Four Typical ELISA Formats The ELISA provides a wealth of information in its simplest formats, but it can also be used in more complex versions to provide enhanced signal, more precise results, or if certain reagents are not available. The four typical ELISA formats are described briefly below. The end result for all the ELISAs is shown in figure 3, a single well, or a series of wells in a multiwall dish, with color intensity varying in proportion to the amount of antigen/analyte in the original sample.

Direct ELISA

An antigen coated to a multiwell plate is detected by an antibody that has been directly conjugated to an enzyme. This can also be reversed, with an antibody coated to the plate and a labeled antigen used for detection, but the second option is less common. This type of ELISA has two main advantages: o It is faster, since fewer steps are required o It is less prone to error, since there are fewer steps and reagents

Indirect ELISA

Antigen coated to a polystyrene multiwell plate is detected in two stages or layers. First an unlabeled primary antibody, which is specific for the antigen, is applied. Next, an enzyme-labeled

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secondary antibody is bound to the first antibody. The secondary antibody is usually an anti-species antibody and is often polyclonal. This method has several advantages: o Increased sensitivity, since more than one labeled antibody is bound per primary antibody o Flexibility, since different primary detection antibodies can be used with a single labeled

secondary antibody o Cost savings, since fewer labeled antibodies are required

Sandwich ELISA

Sandwich ELISAs typically require the use of matched antibody pairs, where each antibody is specific for a different, non-overlapping part (epitope) of the antigen molecule. The first antibody, termed the capture antibody, is coated to the polystyrene plate. Next, the analyte or sample solution is added to the well. A second antibody layer, the detection antibody, follows this step in order to measure the concentration of the analyte. Polyclonals can also be used for capture and/or detection in a sandwich ELISA provided that variability is present in the polyclonal to alow for both capture and detection of the analyte through different epitopes. If the detection antibody is conjugated to an enzyme, then the assay is called a direct sandwich ELISA. If the detection antibody is unlabeled, then a second detection antibody will be needed resulting in an indirect sandwich ELISA. This type of assay has several advantages: o High specificity, since two antibodies are used the antigen/analyte is specifically captured and

detected o Suitable for complex samples, since the antigen does not require purification prior to

measurement o Flexibility and sensitivity, since both direct and indirect detection methods can be used

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Competition or Inhibition ELISA This is the most complex ELISA, and is used to measure the concentration of an antigen (or antibody) in a sample by observing interference in an expected signal output. Hence, it is also referred to as an inhibition ELISA. It can be based upon any of the above ELISA formats, direct, indirect, or sandwich, and as a result it offers maximum flexibility in set up. It is most often used when only one antibody is available to the antigen of interest or when the analyte is small, i.e. a hapten, and cannot be bound by two different antibodies. A simple example of a competitive ELISA is shown in figure 7. In this case samples are added to an ELISA plate containing a known bound antigen. After coating, blocking, and washing steps, unknown samples are added the plate. Detection then follows pretty much as with other ELISA formats. If the antigen in the sample is identical to the plate-adsorbed antigen, then there will be competition for the detection antibody between the bound and free antigen. If there is a high concentration of antigen in the sample, then there will be a significant reduction in signal output of the assay. Conversely, if there is little antigen in the sample, there will be minimal reduction in signal. Therefore, with a competition ELISA, one is actually measuring antigen concentration by noting the extent of the signal reduction. If the detection antibody is labeled, then this would be a direct competition ELISA and if unlabeled, then this would be an indirect competition ELISA. For further examples of competition ELISAs, and a thorough explanation of this technique, please refer to The ELISA Guidebook (Crowther 2001).

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Figure 7. Competition ELISA. Bound and free antigen compete for binding to a

labeled detection antibody.