Introduction:

• DNA Microarray is one of the molecular detection techniques which is a collection of microscopic characteristics (commonly DNA) affixed to a solid surface.
• Also termed as DNA chip or biochip.
• DNA spots can be probed with the target molecules to result either qualitative or quantitative data.
• Microarrays can be characterized on the basis of the nature of the probe, the solid support used and the specific technique used for target detection and/or probe addressing

Approaches of DNA microarrays

Some of the approaches of microarrays are;

1. Printed microarrays
2. In-situ synthesized microarrays
3. High-density bead arrays
4. Electronic microarrays
5. Suspension bead microarrays

1. Printed Microarrays:

• These were first microarrays to be used in research laboratories.
• It is termed as ‘printed microarrays’ because of the printing or spotting of probes on the microarray surface (generally a glass slide).
• The probe spots can be put in either by contact or noncontact printing.
• The direct application of probe solution onto the microarray surface is performed by print pin in contact printing whereas the noncontact printer uses the same technology as the computer printers to eject the droplets of probe solution onto the glass slide.
• Few nanolitres of probe solution per spot is applied to create an array of 100-150mm features, in both the cases of printed microarrays.
• During the course of printing, it is important to control for cross-contamination and conserve the wholeness of the microarray and following hybridization data.
• On account of comparatively large size of the characters, printed microarrays are lower in density(10,000-30,000 characters) than high density bead arrays and in situ-synthesized microarrays but provide more features than suspension bead arrays or electronic microarrays.
• Printed are comparatively simple and economic than in situ  synthesized microarrays. However, the initial step of microarray facilities is expensive and requires separate space where environmental conditions such as humidity, dust, and temperature are strictly controlled.
• A major merit of printed microarrays is flexibility. In clinical     microbiology, printed microarrays are used due to the ability to adjust spotted probes based upon revised annotations or the finding of noble, rising pathogens.
• However, its use is convoluted by the tedious and expensive tasks of observing manufacture reproducibility, performing clinical validation studies, and continually evaluating the quality of downstream data.
• Even if printed microarrays are favourable to user-defined testing, their use in diagnostic microbiology remains confined to specific research applications.

On the basis of nature of the probes, printed microarrays can be classified as:

1. Double stranded DNA microarrays:

• The probes contains of amplicons acquired by PCR using the primers designed either from a familiar genomic sequence, or cDNA or shot-gun library clones.
• The denaturation of the double stranded amplicons takes place either in print buffer or after immobilization, permitting the probes to be available for hybridization.
• The attachment of the amplicons to the glass slide is favoured by the electrostatic interaction of the negative charge bore by the phosphate backbone of the DNA with the positive charge of the coating of the glass slide surface. It can also take place by the formation of UV-cross-linked covalent bonds between the thymidine bases in the DNA and amine groups on slides.
• PCR amplicons are ideally required to have high specificity and yield eliminating contamination, including nonspecific amplification and taints that affect attachment to the microarray surface. But, dsDNA probes have high sensitivity but lacks in specificity. Less specificity can be advantageous when analyzing a genomic sequence rich in natural polymorphisms.
• A great demerit in the production of printed dsDNA microarray is the massive scale of amplicon production and related troubles of quality control, information management, efficiency and preciseness.

2. Oligo-nucleotide microarrays:

• In this type, the spotted probes contains of short, chemically synthesized sequences.
• The probe’s length ranges from 25 to 80bp but it may be as long as 150bp for gene expression microarrays.
• Shorter probe lengths allows less errors during probe synthesis and enables the interrogation of small genomic regions, plus polymorphisms. Specificity is greater when shorter and specific genomic regions are interrogated. With increasing length of the probe, the strength of the hybridization signal and the sensitivity increases.
• In order to improve the hybridization signal strength, the higher concentration of the probes can be applied during printing.
• Despite being easier to produce than dsDNA probes, oligonucleotide probes need to be carefully designed so that all probes acquire similar melting temperatures (within 5⁰ C) and eliminate palindromic sequences.
• The probe’s attachment to the glass slides takes place by the covalent linkage as electrostatic immobilization and cross-linking can result in significant loss of probes during wash steps due to their small size. The coupling of probes to the microarray surface takes place via modified 5′ to 3′ ends on coated slides that provide functional groups (epoxy or aldehyde)

2. In-situ synthesized Arrays:

• In-situ synthesized arrays are immensely-high-density microarrays which use oligonucleotide probes, (out of which Gene chips are most extensively known).
• Contrasting to printed oligonucleotide array, the oligonucleotide probes which is typically 1-2 cm² are synthesized directly on the surface of the microarray.
• As in-situ synthesized probes are usually short(20-25 bp), multiple probes per target are used to enhance statistical accuracy, sensitivity and specificity.
• Normally,11 probes are used per 600 bases being examined.
• The specificity is further increased by use of probe sets. A probe set consists of one perfect-match probe and one mismatch probe that contains a 1bp difference in middle position of probe. Each member of probe set is present in a separate feature, allowing the mismatch probe to behave as a negative control to recognize possible nonspecific cross-hybridization events.
• Recent progresses in Gene Chips consists the use of longer probes, the design of arrays that cross-examine across entire genes or exons, and the execution of multiple self-standing and non-overlapping perfect-match probes in place of classic probe sets.
• Affymetrix Gene Chips classically have >10⁶  characters per microarray depending on the inter-character distance. The probes are produced using semiconductor based photochemical synthesis.
• Synthesis linkers reformed with light-sensitive protecting groups are available on the quartz surface. Thus, the microarray surface is chemically secured from a nucleotide addition.
• When the array surface is exposed to UV light, reactive nucleotides revised with a photolabile protecting group can be joined to increasing oligonucleotide chains.
• Photolithographic masks are used to target specific nucleotides to specific probe sites. Each mask has a designated pattern of windows, which acts as a filter to either transmit or obstruct UV light from specific characters on the chemically preserved microarray surface.
• The areas of the microarray surface that has been exposed to the light will be deprotected and specific nucleotides can be added. The pattern of windows in each mask controls the sequence of nucleotide addition.
• In-situ probe synthesis is thus completed through the cycling of masking, exposure to light, and the addition of either A,T,C or G bases to the growing oligonucleotide.
• Further high density oligonucleotide arrays include those synthesized by Roche NimbleGen and Agilent technologies. These platforms use longer oligonucleotide probes(60-100bp), but NimbleGen and Agilent uses maskless photo-mediated manufacture, and inkjet technology respectively for the synthesis of probes.
• Also, NimbleGen and Agilent platforms allow multi-color hybridization whereas experiments performed with GeneChips are confined to one label.
• Synthesized microarrays relies on commercial synthesis due to complicated nature of chemical synthesis and huge expense required in production and are therefore favourable for user-defined progress.
• The major merits to these systems are the duplicatibility of the production process and the standardization of reagents, instrumentation, and data analysis, all of which are applied to the clinical laboratory.
• Regardless printed or synthesized, oligonucleotide arrays normally permits much cleaner downstream hybridization than amplicon based microarrays.
• Although in-situ synthesized oligonucleotide microarrays are very vigorous systems and have important control measures included, there are recently none with direct diagnostic contagious disease applications that are commercially accessible.

3. High-density bead arrays:

• Bead arrays issues a patterned substrate for the high-density spotting of target nuclei acids alike the printed and in situ-hybridized microarrays.
• Bead arrays rely on 3-mm silica beads that haphazardly self gather onto one of two available substrates: the Sentrix Array Matrix (SAM) or the Sentrix Bead Chip.
• The SAM comprises 96 1.4-mm fibre-optic bundles. Each bundle is a single array containing of 50,000 5-mm light conducting fibers, each of which is chemically engraved to form a microwell for a single bead.
• Upto 1,536 types of bead gather onto each fibre bundle, resulting in 30 beads of each type in the array in case of universal bead array.
• Each SAM permits the inquiry of 96 independent samples.
• Bead chips are more suitable for high density applications such as whole-genome typing.
• Contrast to the known locations of printed and in situ hybridized microarray characters, the beads in Bead arrays haphazardly group to their final location on the array.
• Thus the bead location is mapped which is completed by a decoding process.
• The SAM can be treated using a standard microtiter plate, which makes it manageable to standard automation and high-throughput processing.
• The distance between individual arrays on the 16-sample Bead Chip is similar to that of a standard multichannel pipettor, thereby mediating ease of use.
• Bead arrays can support up to 105 to 106 characters and have built-in repetition.
• Since each manufactured microarray will not be similar, this redundancy is a pivotal experimental control for inter microarray comparative data.
• An additional merit to the peculiarity of each microarray is that altering the bead pattern produces a method to identify spatial bias.
• Although the analysis tools are present for Bead array-specific data analysis, background rectification, and spatial trace recognition have been delaying behind those provided by other microarray manufacturers.
• Bead arrays have been usually applied to DNA methylation studies, gene expression profiling , and SNP genotyping.

4. Electronic microarrays:

• The printed and in situ-synthesized microarrays and Bead arrays explained earlier depend on passive transport for the hybridization of nucleic acids. Dissimilar to it, electronic microarrays utilize active hybridization via electric fields to control nucleic acid transport.
• For the electronic addressing of nucleic acids, microelectronic cartridges use complementary metal oxide semiconductor technology.
• Each Nano Chip cartridge consists of 12 connectors that control 400 individual test sites.
• When a positive current is applied to one or more test sites on the microarray, negatively charged nucleic acids are transported to specific sites, or features.
• The surface of the microarray consists  streptavidin, which forms streptavidin-biotin bonds once electronically addressed biotinylated probes meet their targeted location. The positive current is then eliminated from the active features, and new test sites can be operated by the targeted application of a positive current.
• The microarray is set  for the application of fluorescently labeled target DNA, once the probes have been hybridized at distinct features.
• Usually, target DNA passively hybridizes with the immobilized probes on the microarray but can also be concentrated electronically. Even though addressing the capture probe down first is the most normally used format, amplicon-down and sandwich assays have also been employed.
• Regardless of the addressing format used, if hybridization occurs between the probe and the target DNA, fluorescent reporters will be present at the positive test, which will be detected when the electronic microarray is examined and analysed.
• Electronic microarrays has many merits. For instance, since multiple probes, each with a discrete fluorophore, can be sequentially addressed to the same feature, multiplex detection can be completed at a single test site.
• The flexibility of this platform permits nucleic acids from an individual sample to be hybridized to multiple test sites for the spotting of multiple targets, or nucleic acids from multiple samples can be examined on the same microarray cartridge, decreasing waste.

5. Suspension bead arrays:

•  Dissimilar to the two-dimensional, or planar, arrays explained earlier, suspension bead arrays are three- dimensional arrays which are based on the utilization of microscopic polystyrene spheres (beads) as the solid support and flow cytometry for bead and target spotting.
• Furthermore, they are discrete from the high-density Illumina Bead arrays discussed earlier, in which the beads are placed on fiber- optic strands or silicon slides.
• Suspension-bead-based assays were initially described in 1977 and focused on the detection of antibodies and antigens.
• Multiplexing was initially achieved by using variable-sized microsphere sets for the continuous detection of multiple antibodies. Currently, more vigorous multiplexing is completed using different microsphere sets based on color.
• Red (658-nm radiation) and infra- red (712-nm radiation) fluorochromes are used at different concentrations to fill 5.6-mm microspheres.
• Each bead of the 100-microsphere set has a different red-to-infrared ratio, and thus, each bead has a distinct spectral address.
• Microspheres with a specific spectral address integrated to a specific probe are identical to a character in a planar microarray.
• Once multiple individual microspheres have been incorporated to separate specific probes, a mixture of microspheres can be used to examine extracted and amplified nucleic acids.
• Using a bench-top flow cytometer, the succeeding  detection of a fluorescent reporter that indicates probe-target DNA hybridization is completed.
• An individual file microsphere suspension proceeds by two lasers. A 635-nm laser stimulates the red and infrared fluorochromes infused in the microspheres, which permits the classification of the bead and thus the identity of the probe-target being examined. A 532-nm laser thrills reporter fluorochromes such as R-phycoerythrin and Alexa 532 to determine any hybridization that takes place on the microsphere.
• For nucleic acid detection by suspension bead arrays, several chemistries have been initiated that includes direct DNA hybridization, competitive DNA hybridization, and solution based chemistries with microsphere capture. In direct DNA hybridization, PCR amplicons hybridize directly to probe capture sequences placed on the microspheres.
• Principally, a biotinylated primer used during amplification permits streptavidin–R-phycoerythrin to attach and label hybridized microspheres.
• Competitive DNA hybridization employs unlabeled PCR amplicons and biotinylated competitor oligonucleotides. In difference to the direct hybridization method, competitive DNA hybridization results high fluorescence in the lack of target DNA. When target DNA is available, it binds the tagged competitor DNA, which, in turn, is not present to hybridize to the microsphere, resulting low fluorescence.
• Allele-specific primer extension (ASPE) or target-specific primer extension (TSPE), oligonucleotide ligation assay (OLA), and single-base-chain extension (SBCE) are solution-based chemistries integrated with succeeding microsphere capture.
• By utilizing the natural properties of DNA polymerases and ligases, these chemistries integrate a capture sequence during the solution-based reaction.
• Both ASPE or TSPE and OLA use a capture primer, which consists a unique 5′ sequence succeeded  by a target-specific sequence. In ASPE and/or TSPE, the primer can be extended by DNA polymerase only if target DNA is available to supply the complementary base for the 3′ nucleotide. The tag in ASPE and/or TSPE is facilitated by a biotinylated deoxynucleotide triphosphate. The OLA reaction is ligase dependent. In addition to the capture primer, a biotinylated probe homologous to target DNA is present during an OLA.
• The capture primer and reporter probes can be joined only if target DNA is present in the sample. Used for multiplex SNP detection, SBCE needs independent reactions for each nucleotide query. For every SNP being cross-examined, one probe with a distinct capture sequence is used to assay the possible alleles in separate wells consisting a distinct dideoxynucleoside triphosphate per well. When the capture and target sequences are homologous, a biotinylated dideoxynucleoside triphosphate is integrated, thereby stopping further augmentation.
• The solution-based chemistries explained above all take advantage of universal microspheres with nonspecific capture sequences.
• The first universal sequences used to label microspheres were ZipCode/cZipCode arrest sequences originally used with SBCE in SNP genotyping assays. The 25-bp ZipCode sequences are based on erratic genomic sequences from Mycobacterium tuberculosis.
• A distinct ZipCode sequence is incorporated in the 5′ end of the capture probe used in the chemistries explained above, while microspheres are labeled with the complementary sequence.
• Even though the character density of suspension bead arrays is the least of all the platforms assessed, merits abound that make this platform the most practical for clinical microbiology applications.
• The presence of universal bead sets and their innate flexibility make the progress of user-defined applications practical and relatively economic. Although users must carefully affirm the positive fluorescent threshold for each analyte in the multiplex, user-defined bead-based assays allow experienced users a multitude of clinically relevant applications.
• Significantly, in 2008, Luminex obtained FDA clearance for the first infectious-disease suspension bead array (xTAG RVP), which diagnose 12 respiratory viruses and subtypes. Although analyte-specific reagents (ASRs) also exist, the presence of FDA-cleared products is an important step in getting this technology into less-experienced detective microbiology laboratories.
• However, many built clinical molecular microbiology laboratories depends greatly on real-time PCR, which has less contamination risks.
• In distinction, the opening of postamplification tubes and the following pipetting steps in the workflow of suspension arrays increase the risk for intra- and inter run contamination. Careful deliberation should be taken to control contamination and the reestablishment of postamplification laboratory space in the era of real-time PCR.
• Nevertheless, the relative clarity, strong multiplexing abilities, and cost effectiveness of suspension bead arrays make this platform the most luring for high-throughput nucleic acid diagnosis in clinical contagious disease diagnostics.

References:

• https://www.ncbi.nlm.nih.gov/pmc/articles/PMC3467903/
• http://www.premierbiosoft.com/tech_notes/microarray.html
• https://www.csus.edu/indiv/r/rogersa/bio181/microarrays.pdf
• https://www.csus.edu/indiv/r/rogersa/bio181/microarrays.pdf

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• A copy of DNA generated from messenger RNA (mRNA) with the help of enzyme reverse transcriptase is termed as cDNA.
• A set of cDNA fragments, each of which has been cloned into a separate vector molecule, which constitute a some portion of transcriptome of the organism and stored as a library is known as a cDNA library.

Principle of cDNA library:

• To construct cDNA libraries, DNA copies from mRNA sequences of organism are produced and then they are cloned.
• The term cDNA is given as all the DNA in the library are complementary to the mRNAs and are produced by reverse transcription of mRNAs.
• Most eukaryotic DNA consists of repeated sequences that are not transcribed into mRNA, and in a cDNA library the sequences are not represented.
• It should be remembered that prokaryotes and lower eukaryotes do not contain introns, and cDNA preparation for these species is usually needless.
• Therefore, cDNA libraries are only created from higher eukaryotes.
• For the construction of cDNA library, both the bacterial and bacteriophage DNA can be used as vectors.

Process involved in the construction of cDNA library:

1. Extraction of mRNA from the eukaryotic cell:

• Firstly, the mRNA from the remaining RNAs is collected and purified.
• Many other methods are available for purifying RNA like trizol extrac­tion and column purification.
• By using oligomeric dT nucleotide coated resins, column purification is performed where only the mRNA that has the poly-A tail can bind.
• By using oligomeric dT nucleotide coated resins, column purification is performed where only the mRNA that has the poly-A tail can bind.
• The remaining RNAs are eluted.
• The mRNA is eluted using eluting buffer and also some heat to sepa­rate the mRNA strands from oligo-dT.

2. cDNA construction:

• For the construction of cDNAs, there are several different methods. These are discussed as follows:

i). The RNAse method:

• Principle:
  • By the use of reverse transcriptase, a complementary DNA is synthesized to form an RNA:DNA duplex. Now, the RNA strand is nicked and then replaced by DNA.
• Steps:
• Step I: Annealing:
  • A chemically synthesized oligo-dT primer is annealed to the 3’ polyA-tail of the RNA. The primer is usually 10-15 residues long.
  • In the presence of reverse transcriptase and deoxyribonucleotides, it primes the synthesis of the first DNA strand. This leaves a RNA:DNA duplex.
• Step II: Replacing RNA strand with DNA strand:
  • The RNA strand is replaced by DNA strand by the help of enzyme RNAse H.
  • RNase enzyme removes the RNA from RNA:DNA duplex. The DNA strand which is left behind now acts as a template and the other DNA strand synthesized by the DNA polymerase II.

ii). The self-priming method:

• In this method, the oligo-dT primer is annealed at the polyadenylate tail of the mRNA to prime the first DNA strand synthesis against the mRNA.
• This cDNA, thus formed, tends to fold back on itself temporarily, creating a hairpin loop.
• This results in the second strand’s self-priming.
• This loop must be cleaved with a single-strand-specific nuclease, e.g., SI nuclease, after the synthesis of the second DNA strand to allow insertion into the cloning vector.
• There is a serious drawback to this method.
• At the 5′ end of the clone, cleavage with SI nuclease results in the loss of a certain amount of sequence.

iii). Land et al. strategy:

• The cDNA is tailed with a string of cytidine residues using the enzyme terminal transferase following first-strand synthesis, which is primed with an oligo-dT primer as usual.
• For a synthetic oligo-dG primer, this artificial oligo-dC tail is then used as an annealing site, allowing the second strand to be synthesized.

iv).  Homopolymer tailing:

• The enzyme terminal transferase that can polymerize nucleotides into the 3′-hydroxyl of both DNA and RNA molecules is used in this method.
• In order to generate an RNA: DNA hybrid, the synthesis of the first DNA strand is performed as before.
• In order to add nucleotide tails to the3′ ends of both RNA and DNA strands, then  terminal transferase and a single deoxyribonucleotide is used.
• The consequence of this is that at its3′ end, the DNA strand now has a known sequence. DCTP or dATP are usually used.
• A complementary oligomer (chemically synthesized) can now be annealed and used as a primer to direct the synthesis of the second strand.
• To assist in cloning the resulting double-stranded cDNA, this oligomer (and also the one used for first strand synthesis) can additionally incorporate a restriction site.

v). Rapid amplification of cDNA ends:

• The RACE techniques are split into 3’RACE and 5’RACE, according to the end of the cDNA in which we are interested.
• a.  3’ RACE:
  • Reverse transcriptase synthesis of a first DNA strand is performed using a modified oligo-dT primer in this type of RACE.
  • This primer involves an extension of a particular adaptor sequence followed by an oligo-dT stretch.
  • The first strand synthesis is followed by a second strand synthesis that used a primer internal to the coding sequence of interest.
  • This is accompanied by PCR that uses
    • i. The same internal primer.
    • ii. Sequence of the adaptor (i.e., omitting the oligo-dT). Although it should be possible to use a simple oligo-dT primer in theory instead of the adaptor-oligo-dT and adaptor combination, the low melting temperature can interfere with the subsequent PCR rounds for an oligo-dT primer.
• b. 5’ RACE:
  • The first cDNA strand of this type of RACE is synthesized with re-verse transcriptase and a primer from the coding sequence.
  • It removes the unincorporated primer and tails the cDNA strands with oligo-dA.
  • With an adaptor-oligo-dT primer, a second cDNA strand is then synthesized.
  • The double-stranded molecules resulting from this are then subjected to PCR using
    • i.  A primer nested within the coding region and
    • ii. In the final PCR, a nested primer is used to maximize specificity. Due to the low melting temperature of a basic oligo-dT primer, the adaptor sequence is used in the PCR, as in the 3’RACE above. A variety of kits are commercially available for RACE.

3. cDNA cloning:

a. Linkers:

• In the end, the methods of RNaseH and homopolymer tailing generate a collection of double-stranded, blunt-ended cDNA molecules.
• The vector molecules must now be bound to them.
• This could be achieved by blunt-ended ligation, digestion with the rela-evant enzyme and ligation into the vector, or by adding linkers.

b. Incorporation of restriction sites:

• The homopolymer tailing technique can be adapted by using primers that are adjusted to incorporate restrictions.
• The 3 ‘end of the first cDNA strand, recently synthesized, is tailed with C’s.
• An oligo-dG primer, again preceded by a sail site within the oligonucleotide’s short double-stranded region, is then used for second-strand synthesis.
• The use of an oligonucleotide containing a double-stranded region is necessary in this process.
• Such oligonucleotides are formed by separately synthesizing the two strands and then allowing them to anneal with each other.

c. Homopolymer Tailing of cDNA:

• Another idea is to re-use terminal transferase.
• Treatment with terminal transferase and dCTP of blunt-ended double-stranded cDNA leads to the polymerization of several C residues (typically 20 or so) to 3′ hydroxyl at each end.
• The terminal transferase and dGTP treatment of the vector leads to the inclusion of several G residues on the ends of the vector. It is possible to use dATP and dTTP alternatively.
• It is now possible to anneal the vector and cDNA, and the base-paired region is often so extensive that DNA ligase treatment is unnecessary.
• There may actually be gaps rather than nicks at the edges of the vector insert, but once the recombinant molecules have been inserted into a host, these are repaired by physiological processes.

Advantages of cDNA library:

• There are two major benefits of a cDNA library.
• First, it is enriched with fragments from genes that have been actively transcribed.
• Second, introns do not disrupt the cloned sequences; if the goal is to create a eukaryotic protein in bacteria, introns will pose a problem, since most bacteria have no means of eliminating the introns.

Disadvantages of cDNA library:

• A cDNA library has the drawback that it only includes sequences that are present in mature mRNA.
• There are no introns and any other sequences that are modified during transcription; sequences that are not transcribed into RNA, such as promoters and enhancers, are also not present in a library of cDNA.
• It is also important to remember that only certain gene sequences expressed in the tissue from which the RNA has been isolated constitute the cDNA library.
• In addition, in a cDNA library, the frequency of a specific DNA sequence depends on the abundance of the corresponding mRNA in the given tissue.
• In contrast, in a genomic DNA library, almost all genes are present at the same frequency.

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• Messenger RNA operates as the template for protein synthesis.
• Messenger RNA encodes genetic information from DNA as a transcript and translates the information to a ribosome and helps assemble amino acids in their proper order.
• mRNA is directly transcribed from DNA, whereas in case of eukaryotes, a pre-mRNA (also referred to as the primary transcript) is first transcribed from DNA and then processed to yield mature mRNA.
• Three main regions occur in both prokaryotic and eukaryotic mRNAs.
• 5’ UTR:
  • The 5′ untranslated region (5′ UTR; also called the leader) is a nucleotide sequence at the 5′ end of the mRNA that does not encode any of amino acids.
  • In bacterial mRNA, this region consists of the consensus sequence termed as the Shine-Dalgarno sequence. During translation, Shine-Dalgarno sequence serves as a ribosome binding site. This sequence is found approximately seven nucleotides upstream of the first codon that is translated into the amino acid, also termed as start codon.
  • In its 5′ untranslated region, eukaryotic mRNA has no equivalent consensus sequence.
  • Ribosomes bind to a modified 5′ end of mRNA in eukaryotic cells.
• Protein coding region:
  • The next section of mRNA is the protein-coding region, containing the codons that describe the protein’s amino acid sequence.
  • The protein-coding region starts with a start codon and terminates with a stop codon.
• 3’ UTR:
  • The 3′ untranslated region (3′ UTR; also referred to as a trailer), a nucleotide sequence at the3′ end of the mRNA, is the last mRNA region and not translated into protein.
  • The3′ UTR affects mRNA stability and the translation of the protein-coding sequence of the mRNA.

Post-transcriptional modification in Eukaryotes

• Transcription and translation take place concurrently in bacterial cells; when the 3′ end of an mRNA is undergoing transcription, ribosomes bind near the 5′ end to the Shine-Dalgarno sequence and begin translation.
• Since transcription and translation are coupled, before protein synthesis, bacterial mRNA has little opportunity to be changed.
• In contrast, in eukaryotic cells, transcription and translation are segregated both in time and space.
• In the nucleus, transcription takes place while translation takes place in the cytoplasm; this separation offers a chance to modify eukaryotic RNA before translating it.
• Indeed, after transcription, eukaryotic mRNA is altered extensively.
• Changes are made to the RNA molecule’s 5′ end, the 3′ end, and protein coding portion.
• Following are the examples of Post-transcriptional modification:

1. The 5 ‘Cap Addition:

• One type of eukaryotic pre-mRNA modification is the addition of a structure called a 5 ‘cap at its 5’ end.
• At the 5’ end of the mRNA, the cap consists of an additional nucleotide and methyl groups (CH3) at the base of one or more nucleotides at the 5′ end of the newly inserted nucleotide and the 2′-OH group of sugar.
• After transcription initiation, the insertion of the cap takes place quickly.
• It is possible to represent the 5′ end of pre-mRNA as 5′-pppNpNpN, in which a ribonucleotide is represented by the letter ‘N‘ and a phosphate by ‘p‘.
• One of these phosphate groups is removed shortly after the start of transcription and a guanine nucleotide is added.
• A special 5′-5′ bond connects this guanine nucleotide to the pre mRNA, which is somewhat different from the normal 5′-3′ phosphodiester bond that binds all the other RNA nucleotides.
• To the 5′ end, one or more methyl groups are added.
• The first of these methyl groups is attached to the position 7 of the base of the terminal guanine nucleotide making the base 7-methyl guanine.
• Next, in the second and third nucleotides, a methyl group may be attached to the 2′ position of the sugar.
• Additional methyl groups can rarely be attached to the bases of the second and third nucleotides of  pre-mRNA.

2. The Poly A tail addition:

• The addition of 50 to 250 or more adenine nucleotides at the3′ end, forming a poly(A) tail, is a second kind of modification to eukaryotic mRNA.
• These nucleotides are not encoded in the DNA, but are inserted in a process called polyadenylation following transcription.
• Many RNA polymerase II transcribed eukaryotic genes are transcribed well past the end of the coding sequence; much of the extra material is then cleaved at the3′ end and the poly(A) tail is inserted.
• Sequences both upstream and downstream of the cleavage site are necessary for processing the3′ end of pre-mRNA.
• Generally, downstream of the cleavage site is a sequence rich in uracil nucleotides (or in guanine and uracil nucleotides).
• On many mRNAs, the poly(A) tail confers stability, increasing the time during which the mRNA remains intact and available for translation before cellular enzymes degrade it.
• The poly(A) tail also enhances the ribosome’s attachment to the mRNA.

3. RNA splicing:

• The removal of introns by RNA splicing is the other major type of eukaryotic pre-mRNA modification.
• Before the RNA moves to the cytoplasm, this modification takes place in the nucleus.
• The presence of three sequences in the intron is required for splicing.
• One end of the intron is referred to as the5′ splice site, and the other end is the 3′ splice site; short consensus sequences exist on these splice sites.
• Most introns begin with GU in pre-mRNAs and end with AG.
• The third sequence that is necessary for splicing is present at the branch point, which is an adenine nucleotide that is situated 18-40 nucleotides upstream of the  3′ splice site.
• Splicing occurs within a large structure called the spliceosome, which is one of the largest and most complex of all molecular complexes.
• Five RNA molecules (U1, U2, U4, U5, and U6) and almost 300 proteins form the spliceosome.
• Small nuclear RNAs (snRNAs) ranging in length from 107 to 210 nucleotides are the RNA components; these snRNAs are associated with proteins to form small particles of ribonucleoprotein.

Process of RNA splicing:

• An intron is between an upstream exon (exon1) and a downstream exon (exon 2) before splicing takes place.
• In two distinct stages, pre-mRNA is spliced.
• The pre-mRNA is cut at the 5 ‘splice site in the first stage of splicing.
• This cut frees exon 1 from the intron, and the intron’s 5′ end connects to the branch point; that is, the intron folds back on itself, creating a structure called a lariat.
• In this reaction, via a trans-esterification reaction, the guanine nucleotide in the consensus sequence at the 5′ splice site binds with the adenine nucleotide at the branch stage.
• To the cytoplasm, where it is translated, the mature mRNA consisting of the exons spliced together is exported.
• A cut is made at the3′ splice site in the second step of RNA splicing and, simultaneously, the3′ end of exon 1 is covalently connected (spliced) to the5′ end of exon 2.
• It releases the intron as a lariat.
• When the bond splits at the branch stage, the intron becomes linear and is then quickly degraded by nuclear enzymes.
• To the cytoplasm, where it is translated, the mature mRNA consisting of the exons spliced together is exported.

Alternative processing pathways for RNA splicing:

• In order to generate alternative forms of mRNA, a single pre-mRNA is processed in various ways, resulting in the development of various proteins from the same DNA sequence.
• Alternative splicing, in which the same pre-mRNA can be spliced in more than one way to generate multiple mRNAs that are translated into different amino acid sequences and thus different proteins, is one form of alternative processing.
• Another method of alternative processing involves the use of several 3′ cleavage sites, where the pre-mRNA comprises two or more potential cleavage and polyadenylation sites.
• In the same pre-mRNA transcript, both alternative splicing and multiple 3′ cleavage sites can exist.
• In multicellular eukaryotes, alternative processing of pre-mRNAs is common.
• Researchers predict, that more than 90% of all human genes undergo alternate splicing.
• The type of splicing also varies between human tissues; compared to other tissues, the human brain and liver tissues have more alternatively spliced RNA.

RNA editing:

• The coding sequence of an mRNA molecule is altered after transcription in RNA editing, so that the protein has an amino acid sequence that varies from that of the gene encoded.
• There were substitutions in some of the mRNA nucleotides in some nuclear genes in mammalian cells and in some mitochondrial genes in plant cells.
• More extensive RNA editing for certain mitochondrial genes in trypanosome parasites has been found in the mRNA.
• More than 60 percent of the sequence is determined by RNA editing in some of these organisms’ mRNAs.
• In RNA sequences, a variety of mechanisms can bring about changes.
• Molecules called guide RNAs (gRNAs) play a key role in certain situations.
• gRNAs consist of sequences that are partly complementary to pre-edited RNA segments.
•  In these sequences, the two molecules goes through base pairing.
• The mRNA undergoes cleavage after the mRNA is anchored to the gRNA and nucleotides are inserted, removed or altered according to the gRNA template given.
• Enzymes bring about the conversion of the base in other cases.
• For example, in humans, a gene is transcribed into mRNA that encodes a lipid-transporting polypeptide called apolipoprotein-B100, which is synthesized in liver cells and has 4563 amino acids.
• By editing the apolipoprotein-B100 mRNA, a truncated version of the protein called apolipoprotein-B48 with only 2153 amino acids is synthesized in intestinal cells.
• A cytosine base is deaminated by an enzyme in this editing, transforming it into uracil.
• This conversion converts a codon that specifies the glutamine amino acid into a stop codon that terminates translation prematurely, resulting in the protein being shortened.

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• Complementary DNA (cDNA) cloning is termed for the gene cloning (cloning of DNA fragments) obtained from cDNA.
• The principle of cDNA cloning is that it involves the copying of mRNA transcripts into DNA, which are then inserted into bacterial plasmids and then placed into bacteria by transformation.
• At this stage, it should be clear that mRNA used for cDNA preparation is a processed transcript and not the original one transcribed from DNA.
• In-order to clone a DNA sequence that codes for a required gene product, the gene should be removed from the organism and cloned it in the vector molecule.
• A gene library is a random collection of cloned fragments in an appropriate vector that particularly consists of all the genetic information about that species.
• There are two methods for the formation of gene libraries. They are:
  • Complementary DNA (cDNA)
  • Genomic DNA libraries

Steps of cDNA cloning:

1. Isolation of mRNA
2. Synthesis of first strand of cDNA
3. Synthesis of second strand of cDNA
4. Cloning of cDNA
5. Introduction to host cells
6. Clone selection

1. Isolation of mRNA:

• A crude extract of the tissue with the gene of interest is prepared.
• The extract must be free from proteins, polysaccharides and all other contaminants.
• The technique of oligo-deoxythymine (oligo-dT) cellulose chromatography is used for the further purification of many eukaryotic mRNAs from the total or polysomal fraction.
• mRNAs consist of poly A (adenosine residues) tail at their 3′ end.
• Under favourable conditions, this tail will bind to a string of thymidine residues immobilized on cellulose and then poly (A)⁺ fraction can be eluted.
• Two or three passages of the poly (A)⁺ fraction through such a column produces a fraction highly enriched for mRNA.
• This fraction includes different mRNA sequences, however certain techniques can be employed for extracting a particular mRNA species.
• After the preparation of the fraction, it is essential to confirm if the extracted mRNA consists of the sequence of interest.
• It is performed by translation of mRNA in vitro and identification of suitable polypeptides in the products obtained.

2. Synthesis of first strand of cDNA:

• Reverse transcriptase is a RNA dependent DNA polymerase which is used to copy the mRNA fraction into the first strand of DNA.
• This enzyme, like all other DNA polymerases, can only add residues at the 3′-OH group of an existing primer, which is base paired with the template.
• The most commonly used primer is oligo-dT for cloning of cDNAs.
• Oligo-dT primer is 12-18 nucleotides in length, that binds to the poly (A) tract at the 3′ end of mRNA molecules.
• The RNA strand of the resulting RNA-DNA hybrid is destroyed prior to second strand synthesis through alkaline hydrolysis.

3. Synthesis of second strand of cDNA:

• The second strand of cDNA can be synthesized by two techniques. They are:
• i. Self-priming cDNA:
  • In Self-priming, the mRNA hybrid obtained is denaturated for the synthesis of  second strand on the single strand of cDNA by the klenow fragment of DNA polymerase I.
  • The transitory hairpin structure at the 3′ end of single-stranded DNA can be used to prime the synthesis of second strand of cDNA by the klenow fragment of Escherichia coli DNA polymerase I.
  • Single-strand specific S1 nuclease digests the hairpin loop and any single-stranded overhung at the other end.
  • The ultimate product is a population of double-stranded, blunt-ended DNA molecules complementray to the original mRNA fraction.
• ii. Replacement synthesis:
  • In this method, the cDNA:mRNA hybrid works as a template for a nick translation reaction.
  • In the mRNA strand of the hybrid, RNase H produces nicks and gaps, creating a series of RNA primers.
  • These RNA primers are used by E. coli DNA polymerase I during the synthesis of second strand of cDNA.
  • The advantages of this technique are:
    • –  very efficient
    • –  can be performed directly using the products of the first strand reaction
    • – eliminates the need to use nuclease S1 to cleave the single-stranded hairpin loop in the double stranded cDNA.

4. Cloning of cDNA:

• The most frequently used technique for cloning cDNAs involves the addition of complementary homopolymeric tracts to double stranded cDNA and to the plasmid vector.
• To the cDNA, strings of cytosine residues are added using the enzyme terminal transferase to form oligo-dC tails on the 3′ ends.
• Likewise, a plasmid is cut open at a unique restriction endonuclease site and tailed with oligo-dG.
• Now, the vector and the double stranded cDNA are joined by hydrogen bonding between the complementary homopolymers.
• It results in the formation of open circular hybrid molecules capable of transforming E. coli.

5. Introduction to host cells:

• For the transforming of bacteria, the recombinant plasmids are used, usually the E. coli K-12 strain.
• The uptake of plasmid molecules from the surrounding medium is performed by E. coli cells treated with calcium chloride.
• Any gaps in the recombinant plasmid will be repaired by the host cells.
• The transformed bacteria can be isolated from non-transformed ones on the basis of antibiotic resistance.
• Majority of cloning plasmids contain two antibiotic resistance genes, one of which is destroyed during cloning.
• For instance, in the case of pBR322, cloning into unique PstI site destroys ampicillin resistance but leaves tetracycline resistance intact.
• Bacteria transformed with a recombinant plasmid will be sensitive to ampicillin but resistant to tetracycline.

6. Clone selection:

• The antibiotic resistance selection already performed has recognized which clones carry a recombinant plasmid, however there will be thousands of various inserts.
• The cloning process generally commences with a whole population of mRNA sequences.
• Selection of clones carrying the sequence of interest is the tough job.
• If the gene is expressed, then the simplest selection is to screen for the presence of the protein.
• It can be screened either by bacterial phenotype it produces or by the protein detection methods usually based on immunological or enzymological techniques.
• If the protein is not expressed, then other methods such as nucleic acid hybridization are used.
• Identification of the gene is discussed after the genomic DNA cloning.

cDNA cloning: Principle and steps involved in cDNA cloning

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• Analytical devices that consists a combination of biological detecting elements like sensor system and a transducer is termed as biosensor.
• Biosensors can be defined as self-sufficient integrated devices that has capacity to provide specific qualitative or semi-quantitative analytical information using a biological recognition element which is in direct-spatial contact with a transductional element.
• In simple words, biosensors are analytical devices that detects changes in biological processes and transform the biological data into electrical signal.
• The main features of biosensors are:
  • Stability
  • Economical
  • Sensitivity
  • Reproducibility

Components of biosensor:

• The block diagram of the biosensor consists of three segments namely, sensor, transducer, and electrical circuit.
  • i. Sensor or detector: The first segment is the sensor or detector which is a biological component. it is a biochemical receptor. It interacts with the analyte and signal the change in its composition as electrical signal.
  • ii. Transducer: The second segment is the transducer and it is a physical component which amplifies the biochemical signal received from detector, alters the resulting signal into electrical and displays in an attainable way.
  • iii. Electrical circuit: It is the associated part which consists of Signal Conditioning Unit, a Processor or Micro-controller and a Display Unit.

Principle of Biosensors:

• Biosensors works on the principle of signal transduction and biorecognition of element.
• All the biological materials including-enzyme, antibody, nucleic acid, hormone, organelle or whole cell can be used as sensor or detector in a device. But the desired bio-receptor is usually a specific deactivated enzyme.
• The deactivated enzyme is placed in proximity to the transducer.
• The tested analyte links to the specific enzyme (bio-receptor) and inducing a change in biochemical property of enzyme. The change in in turn gives an electronic response through an electroenzymatic approach.
• Electroenzymatic process is the chemical process of converting the enzymes into corresponding electrical signals with the aid of transducer.
• Now, the outcome from transducer i.e. electrical signal is a direct representation of the biological material (i.e. analyte and enzyme in this case) being measured.
• The electrical signal is usually converted into physical display for its proper analysis and representation.

Working principle of biosensors:

• The union of biological sensitive element and a transducer is responsible to convert the biological material into a corresponding electrical response in form of signal.
• The output of the transducer will be either current or voltage relying on the type of enzyme.
• If the output is voltage, then it is fine. But if the output is current, then this current needs to be converted into equivalent voltage (using an Op-Amp based current to voltage converter) before proceeding further.
• The output voltage signal is generally very low in amplitude and is superimposed on a high frequency noise signal.
• Thus, the signal is amplified (using an Op-Amp based Amplifier) and then it is  passed through a Low Pass RC Filter.
• Signal Processing Unit or a Signal Conditioning Unit is accountable for performing the this process of amplifying and filtering the signal .
• The output of the signal processing unit is termed as an analog signal. This output is equivalent to the biological quantity being measured.
• The analog signal can be exhibited directly on an LCD display but usually, this analog signal is passed to a Microcontroller, where the analog signal is converted into digital signal. This is done since it is easy to analyse, process or store a digital signal.

Types:

• On the basis of sensor device as well as the biological material the biosensors are classified as:
  • 1. Electrochemical biosensors
  • 2. Calorimetric/Thermal detection biosensors
  • 3. Optical biosensors
  • 4. Piezo-electric biosensors
  • 5. Resonant biosensors

1. Electrochemical biosensors:

• Generally, electrochemical biosensor works on the principle that many enzyme catalysis reactions consumes or generates ions or electrons causing some change in electrical properties of the solution which can be detected and used as a measuring parameter.
• For example some biological compounds such as glucose, urea, cholesterol, etc.) are not electroactive, so the combination of reactions by this biosensor produce an electroactive element. This electroactive element results in change of current intensity which is proportional to the concentration of analyte.
• An electrochemical biosensor uses an electrochemical cell with electrodes of different dimension and modifications.
• Three kinds of electrodes are generally used-
  • Working electrode
  • Reference electrode
  • Counter or Auxilary electrode
• It is the working electrode where reaction occurs between electrode substrate and analyte.

Types of electrochemical biosensors

• Electrochemical biosensors are classified into three types:
  • Amperometric Biosensors
  • Potentiometric Biosensors
  • Conductimetric Biosensors
• 1. Amperometric Biosensors
  • The Bioelectrochemical reaction in this biosensors generate measurable amount of current which is directly proportional to the substrate concentration.
  • The first generation amperometric biosensors use the Clark oxygen electrode which determines the reduction of O2 present in the analyte solution.
  • Determination of glucose using glucose oxidase enzyme is a redox reaction which is an example of Amperometric biosensors.
  • This first generation biosensors depend on the dissolved O2 to measure the concentration analyte. However, as modification in second generation biosensors, a mediators is being used.
  • This mediators transfer the electrons produced by the bioelectrochemical reaction directly to the electrode instead of reducing O2 dissolved in analyte solution.
  • Nowadays, the electrodes remove the electrons without the aid of mediators and are coated with electrically conducting organic salts.
• 2. Potentiometric biosensors:
  • Potentiometric biosensors use the ion-selective electrodes to convert the biological reaction to electronic response.
  • Most commonly used electrodes are pH meter glass electrodes (for cations glass pH electrodes coated with a gas selective membrane for CO2, NH or H₂S.) or solid state electrodes.
  • Biosensors detects and measures the ions or electrons generated in many reactions, very weak buffer solutions are used in this case.
  • Gas sensing electrodes detect and measure the amount of gas produced.
• 3. Conductimetric biosensors:
  • These biosensors measure electrical conductance/ resistance of the solution.
  • Conductance measurement have comparatively low sensitivity.
  • Electrical field is generated by use of sinusoidal (ac) voltage which serves in reducing unwanted effects such as:
    • Faradaic processes
    • Double layer charging
    • Concentration polarization

2. Calorimetric/Thermal detection biosensors:

• Most of the enzyme catalysed reactions are exothermic in nature.
• Calorimetric biosensors measure the change in temperature of analyte solution following enzyme action and interpret it in terms of analyte concentration in the solution.
• The analyte solution is passed through a small packed bed column consisting immobilized enzyme.
• The temperature of the solution is measured just before the entry of the solution into the column, and just as it leaves the column using separate thermistors.
• It is the most usually applicable type of biosensor and can also be used for turbid and colourful solutions.
• There are demerits such as:
  • The biggest demerit is to maintain the temperature of the sample stream say + or -0.01°C.
  • Low range and sensitivity.

3. Optical biosensors:

• Both catalytic and affinity reactions are measured by this biosensor.
• The products generated during the catalytic reactions cause a change in fluorescence that is measured by the biosensor.
• In other way, biosensors measure the change induced in the intrinsic optical properties of the biosensor surface due to loading on it of di-electric molecules like protein.
• A most advanced biosensor involving luminescence uses luciferase enzyme for detection of bacteria in food or clinical samples.
• In the presence of O2, luciferase takes up the ATP released from the lysis of bacteria to produce light which is detected and measured by biosensor.

4. Piezo-electric biosensors:

• In these biosensors, the surface is coated with antibodies which binds to the complementary antigen present in the sample solution.
• This results in increased mass which decreases their vibrational frequency, this alteration/change is used to determine the amount of antigen present in the sample solution.

4. Resonant biosensors:

• The vibrations of the electron cloud in a molecule is termed as resonant biosensors.
• These plasmons oscillate at a particular frequency characteristic of the material.
• The oscillations in surface plasmons are confined to the surface of the material.
• Generally, gold or silver surfaces are preferred for the SPR based biosensors.
• When electromagnetic radiation falls on the metal surface, at a particular angle of incidence, the frequency of the electromagnetic radiation matches the frequency of vibrations resulting in resonance.
• The resonant angle depends on the refractive index of the medium.
• The refractive index in turn is determined by the local mass density on the metal surface.
• If the surface of the metal film is modified with the antibody/receptor i.e. capture molecule, then specific binding occurs between the capture molecule on addition of the sample and its ligand leading to an alteration in mass and hence change in resonant angle.
• These biosensors are employed to understand the functional aspects of human immune deficiency virus (HIV) both qualitatively and quantitatively.
• The major merits of these biosensors are rapid measurements and relatively high sensitivity,
• The major demerit is that it cannot be used to detect and measure the turbid and coloured solutions. In few cases, ligands may interfere with the binding.

Biosensors: Components, Working principle and Types

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• Also termed as DNA chips, gene chips, DNA arrays, gene arrays and biochips.
• Biochips are latest generation of biosensors developed by use of DNA probes.
• DNA microarray is one of the molecular detection techniques which is a collection of microscopic characteristics (commonly DNA) affixed to a solid surface.
• DNA microarrays are solid supports usually made up of glass or silicon upon which DNA is attached in an organized pre-arranged grid design.
• Each spot of DNA, termed as probe, signifies a single gene.
• DNA microarrays can examine the expression of tens of thousands of genes concurrently.
• There are 2 types of DNA microarray i.e. cDNA based microarray and oligonucleotide based microarray.

Principle of DNA microarray:

• DNA microarray technology was originated from Southern blotting, in which fragmented DNA is attached to a substrate and then probed with a known DNA sequence.
• DNA microarray is based on principle of hybridization between the nucleic acid strands.
• Complementary nucleic acid sequences have the characteristic to specifically pair to each other by the formation of hydrogen bonds between complementary nucleotide base pairs.
•   Unknown sample of DNA sequence is termed as sample or target and the known sequence of DNA molecule is called as probe.
• Fluorescent dyes are used for labelling the samples and at least 2 samples are hybridized to the chip.
• A large number of complementary base pairs in nucleotide sequence is suggestive of tighter non-covalent bonding between the two strands.
• Following the washing off of non-specific bonding sequences, only strongly paired strands will stay hybridized.
• Thus, the fluorescent labeled target sequences that pairs to the probe releases a signal that relies on the strength of the hybridization detected by the number of paired bases, hybridization conditions, and washing after hybridization.
• DNA microarrays employs relative quantization in which the comparison of same character is done under two different conditions and the identification of that character is known by its position.
• After completion of the hybridization, the surface of chip can be examined both qualitatively and quantitatively by use of autoradiography, laser scanning, fluorescence detection device, enzyme detection system.
• The presence of one genomic or cDNA sequence in 1,00,000 or more can be screened in a single hybridization by using DNA microarray.

Types of DNA microarray:

1. cDNA based microarrays
2. Oligonucleotide based microarrays

cDNA based microarrays:

• cDNA is used for the preparation of chips.
• cDNAs are amplified by PCR.
• It is a high throughput technique.
• It is highly parallel RNA expression assay technique that allows quantitative analysis of RNAs transcribed from both known and unknown genes.

Oligonucleotide based microarrays:

• In this type, the spotted probes contains of short, chemically synthesized sequences, 20-25 mers/gene.
• Shorter probe lengths allows less errors during probe synthesis and enables the interrogation of small genomic regions, plus polymorphisms
• Despite being easier to produce than dsDNA probes, oligonucleotide probes need to be carefully designed so that all probes acquire similar melting temperatures (within 5⁰ c) and eliminate palindromic sequences.
• The probe’s attachment to the glass slides takes place by the covalent linkage as electrostatic immobilization and cross-linking can result in significant loss of probes during wash steps due to their small size.
• The coupling of probes to the microarray surface takes place via modified 5′ to 3′ ends on coated slides that provide functional groups (epoxy or aldehyde)

Requirements of DNA microarray:

• DNA chip
• Fluorescent dyes
• Fluorescent labelled target/sample
• Probes
• Scanner

Steps involved in cDNA based microarray:

1. Sample collection
2. Isolation of mRNA
3. Creation of labeled cDNA
4. Hybridization
5. Collection and analysis

Sample collection: 

• A sample can be any cell/tissue that we desire to conduct our study on.
• Generally, 2 types of samples are collected, i.e. healthy and infected cells, for comparing and obtaining the results.

Isolation of mRNA:

• The extraction of RNA from a sample is performed by using a column or solvent like phenol-chloroform.
• mRNA is isolated from the extracted RNA leaving behind rRNA and tRNA.
• As mRNA has a poly-A tail, column beads with poly-T tails are employed to bind mRNA.
• Following the extraction, buffer is used to rinse the column inorder to isolate mRNA from the beads.

Creation of labeled cDNA:

• Reverse transcription of mRNA yields cDNA.
• Both the samples are then integrated with different fluorescent dyes for the production of fluorescent cDNA strands which allows to differentiate the sample category of the cDNAs.

Hybridization:

• The labeled cDNAs from both the samples are placed on the DNA microarray which permits the hybridization of each cDNA to its complementary strand.
• Then they are thoroughly washed to remove unpaired sequences.

Collection and analysis:

• Microarray scanner is used to collect the data.
• The scanner contains a laser, a computer and a camera. The laser is responsible for exciting the fluorescence of the cDNA, generating signals.
• The camera records the images produced at the time laser scans the array.
• Then computer stores the data and yields results instantly. The data are now analyzed.
• The distinct intensity of the colors for each spot determines the character of the gene in that particular spot.

Applications of DNA microarray technique:

• Drug discovery
• Study of functional genomics
• DNA sequencing
• Gene expression profiling
• Study of proteomics
• Diagnostics and genetic engineering
• Toxicological researches
• Pharmacogenomics and theranostics

DNA Microarray: Principle, Types and steps involved in cDNA microarrays

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Difference between Live and Killed vaccine

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• Vaccine is a biological preparation that improves immunity to a particular disease. They are molecules, usually but not necessarily proteins, that elicits an immune response, thereby providing protective immunity against a potential pathogen.
• Vaccine is used to boost the body’s immune system and prevent the serious life threatening diseases.
• Vaccine can be prepared against bacteria or even eukaryotic protozoans, however most successful vaccines have been developed against viruses.
• The first human vaccine was smallpox vaccine, using cowpox as vaccine.
• Rabies was the first virus attenuated in a lab to create a vaccine for humans.

Characteristics of Vaccine:

Following are the properties that an ideal vaccine should possesses;

1. Safe: Vaccine must be safe and must not itself causes illness or death
2. Protective: vaccine must protect against illness resulting from exposure to live pathogen
3. Sustained protection: protection against illness must last for years
4. Induce neutralizing antibodies
5. Induce protective T cells
6. Low cost
7. Biological stability
8. Ease of administration
9. No side effect or Very few side effects

Types of vaccines

1, Live attenuated vaccines:

• Live attenuated vaccines contain living microorganisms that has been weakened in the lab so it can’t cause disease. These vaccine evoke immune system of the host preventing form the diseases.
• Live attenuated vaccines are relatively easy to create for certain viruses.
• Vaccines against measles, mumps, and chickenpox, for example, are made by this method.
• Live attenuated vaccines are more difficult to create for bacteria. Bacteria have thousands of genes and thus are much harder to control. However, this approach has been used to create a vaccine against Vibrio cholerae.
• Live attenuated vaccine use whole organism as vaccine and are prepared from attenuated strains that almost or completely devoid of pathogenicity but are capable of inducing a protective immune response. They multiply in the human host and provide continuous antigenic stimulation over a period of time.
• Examples: BCG, Typhoid vaccine, Measles vaccine, mumps vaccine, Sabin’s polio vaccine, VAR vaccine, Yellow fever vaccine, Rota virus vaccine etc

2. Killed or Inactivated vaccines: 

• Disease-causing microorganisms are killed with chemicals, heat, or radiation. Such vaccines are more stable and safer than live vaccines because the dead microorganisms can’t mutate back to their disease-causing state.
• There are the easiest preparations to use. Such vaccines are simply inactivated or killed microorganisms.
• Preparation of killed vaccine may take the route of heat or chemicals. The chemicals used includes formaldehyde or beta-propiolactone. The traditional agent for inactivation of virus is formalin.
• Excessive treatment can destroy immunogenicity whereas insufficient treatment can leave infectious microorganisms capable of causing disease.
• The inactivated vaccines usually don’t require refrigeration, and they can be easily stored and transported in a freeze-dried form, which makes them accessible to people in developing countries.
• Example: Salk polio vaccine, Anthrax vaccine, Cholera vaccine, Purtusis vaccine, Plague vaccine, Influenza vaccine, Hepatitis A vaccine, Rabies vaccine, Rubella vaccine etc

3. Subunit vaccines:

• Instead of the entire microorganisms, subunit vaccines include only the antigens that best stimulate the immune system and used in vaccine preparation.
• Vaccine that consists of specific, purified macromolecules derived from pathogen are known as subunit vaccine.
• The general forms of such vaccine are in current use: Purified capsular polysaccharides, inactivated exotoxin (Toxoid), recombinant microbial antigen, synthetic peptide.

i. Purified capsular polysaccharide vaccine:

• The virulence of some pathogenic bacteria depends primarily on the anti-phagocytic property of their hydrophobic polysaccharide capsule.
• These are generally conjugate vaccine.
• In some gran negative bacteria, LPS is the outermost covering which protect the bacteria from binding with the antibody. So that the immature immune systems of infants and younger children can’t recognize or respond to them. For this conjugate vaccine is used.
• Examples: Hib vaccine (The vaccine that protects against Haemophilus influenzae type B (Hib) is a conjugate vaccine), Vaccine for Streptococcus pneumoniae, vaccine for Neisseria meningitides

ii. Toxoid vaccines:

• For some bacteria that secrete toxins, or harmful chemicals, a toxoid vaccine is made.
• The toxins are inactivated by treating with formalin, such detoxified toxin is known as toxoid, which is used as vaccine.
• Vaccination with toxoid induces anti-toxoid antibodies, which are capable of binding the toxin and neutralizing its effect.
• Conditions for the production of toxoid vaccines must be closely controlled to achieve detoxification without excessive modification of the epitope structure. Sufficient quantities of the purified toxins is prepared by cloning the exotoxin genes and then expressing them in easily grown host cells, purified and subsequently inactivated.
• Vaccines against diphtheria and tetanus are examples of toxoid vaccines.

iii. Recombinant vaccine:

• Recombinant vector vaccines are experimental vaccines similar to DNA vaccines, but they use an attenuated virus or bacterium to introduce microbial DNA to cells of the body.
• Attenuated bacteria also can be used as vectors.
• The gene coding for immunogenic protein is inserted into plasmid vector and then transformed it into suitable host cell such as bacteria, yeast, mammal cell etc.
• In this case, the inserted genetic material causes the bacteria to display the antigens of other microbes on its surface.
• In effect, the harmless bacterium mimics a harmful microbe, provoking an immune response.
• Recombinant hepatitis B vaccine is the only recombinant vaccine licensed at present.

iv. Synthetic peptide vaccine:

• The development of synthetic peptides that might be useful as vaccines depends on the identification of immunogenic sites.
• The best example is Foot and mouth disease where protection was achieved by immunizing animals with a linear sequences of 20 aminoacids.
• Synthetic peptide vaccine would have many advantages. They are stable and relatively cheap to manufacture. Furthermore, less quality assurance is required.
• Synthetic peptides so not readily stimulate t cells. It was generally assume that, because of their small size, peptides would behave like haptens and would therefore require coupling to a protein carrier which is recognized by T cells.
• It is now known that synthetic peptides can be highly immunogenic in their free form provided they contain, in addition to the B cells epitope, T cell epitope recognized by T-helper cells. Such T cell epitope can be provided by carrier protein molecules, foreign antigens or within the synthetic peptide molecule itself.
• Synthetic peptide vaccine is not applicable for all viruses. For example, it is not applicable for Polio virus because important antigenic sites were made up of 2 or more different viral capsid protein.

4. DNA vaccines:

• DNA vaccine is DNA sequence used as vaccine.
• This DNA sequence codes for antigenic protein of pathogen.
• When the genes for a microbe’s antigens are introduced into the body, some cells will take up that DNA. The DNA then instructs those cells to make the antigen molecules. The cells secrete the antigens and display them on their surfaces. In other words, the body’s own cells become vaccine-making factories, creating the antigens necessary to stimulate the immune system.
• As this DNA inserted into cells it is translated to form antigenic protein. As this protein is foreign to cells, immune response raised against this protein. In this way, DNA vaccine provide immunity against that pathogen.
• Recently, encouraging results were reported for DNA vaccines whereas DNA coding for the foreign antigen is directly injected into the animal so that foreign antigen is directly produced by the host cells.
• In theory these vaccines would be extremely safe and devoid of side effects since the foreign antigens would be directly produced by the host animal.
• In addition, DNA is relatively inexpensive and easier to produce than conventional vaccines and thus this technology may one day increase the availability of vaccines to developing countries.
• The time for development of DNA vaccine is relatively short which enable timely immunization against emerging infectious diseases.
• Also, DNA vaccines can theoretically result in more long term production of an antigenic protein when introduced into a relatively non dividing tissue such as muscles.
• Examples,  DNA Vaccine Against West Nile Virus, Influenza and Herpes virus

Vaccine: Characteristics and types of vaccine

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Principle:

• Western blotting technique is used for identification of particular protein from the mixture of protein.
• In this method labelled antibody against particular protein is used identify the desired protein, so it is a specific test. Western blotting is also known as immunoblotting because it uses antibodies to detect the protein.

Procedure/Steps:

1. Extraction of protein
2. Gel electrophoresis: SDS PAGE
3. Blotting: electrical or capillary blotting
4. Blocking: BSA
5. Treatment with primary antibody
6. Treatment with secondary antibody( enzyme labelled anti Ab)
7. Treatment with specific substrate; if enzyme is alkaline phosphatase, substrate is p-nitro phenyl phosphate which give color.

Step I: Extraction of Protein

• Cell lysate is most common sample for western blotting.
• Protein is extracted from cell by mechanical or chemical lysis of cell. This step is also known as tissue preparation.
• To prevent denaturing of protein protease inhibitor is used.
• The concentration of protein is determined by spectroscopy.
• When sufficient amount of protein sample is obtained, it is diluted in loading buffer containing glycerol which helps to sink the sample in well.
• Tracking dye (bromothymol blue) is also added in sample to monitor the movement of proteins.

Step II: Gel electrophoresis

• The sample is loaded in well of SDS-PAGE Sodium dodecyl sulfate- poly-acrylamide gel electrophoresis.
• The proteins are separated on the basis of electric charge, isoelectric point, molecular weight, or combination of these all.
• The small size protein moves faster than large size protein.
• Protein are negatively charged, so they move toward positive (anode) pole as electric current is applied.

Step III: Blotting

• The nitrocellulose membrane is placed on the gel. The separated protein from gel get transferred to nitrocellulose paper by capillary action. This type of blotting is time consuming and may take 1-2 days
• For fast and more efficient transfer of desired protein from the gel to nitrocellulose paper electro-blotting can be used.
• In electro-blotting nitrocellulose membrane is sandwich between gel and cassette of filter paper and then electric current is passed through the gel causing transfer of protein to the membrane.

Step IV: Blocking

• Blocking is very important step in western blotting.
• Antibodies are also protein so they are likely to bind the nitrocellulose paper. So before adding the primary antibody the membrane is non-specifically saturated or masked by using casein or Bovine serum albumin (BSA).

Step V: Treatment with Primary Antibody

• The primary antibody (1° Ab) is specific to desired protein so it form Ag-Ab complex

Step VI: Treatment with secondary antibody

• The secondary antibody is enzyme labelled. For eg. alkaline phosphatase or Horseradish peroxidase (HRP) is labelled with secondary antibody.
• Secondary antibody (2° Ab) is antibody against primary antibody (anti-antibody) so it can bind with Ag-Ab complex.

Step VII: Treatment with suitable substrate

• To visualize the enzyme action, the reaction mixture is incubated with specific substrate.
• The enzyme convert the substrate to give visible colored product, so band of color can be visualized in the membrane.
• Western blotting is also a quantitative test to determine the amount of protein in sample.

Application:

1. To determine the size and amount of protein in given sample.
2. Disease diagnosis: detects antibody against virus or bacteria in serum.
3. Western blotting technique is the confirmatory test for HIV. It detects anti HIV antibody in patient’s serum.
4. Useful to detect defective proteins. For eg Prions disease.
5. Definitive test for Creutzfeldt-Jacob disease, Lyme disease, Hepatitis B and Herpes

Western blotting technique: principle, procedure and application

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Principle:

• Southern blotting is an example of RFLP (restriction fragment length polymorphism). It was developed by Edward M. Southern (1975). Southern blotting is a hybridization technique for identification of particular size of DNA from the mixture of other similar molecules. This technique is based on the principle of separation of DNA fragments by gel electrophoresis and identified by labelled probe hybridization.
• Basically, the DNA fragments are separated on the basis of size and charge during electrophoresis. Separated DNA fragments after transferring on nylon membrane, the desired DNA is detected using specific DNA probe that is complementary to the desired DNA.
• A hybridization probe is a short (100-500bp), single stranded DNA. The probes are labeled with a marker so that they can be detected after hybridization.

Procedure/ Steps

1. Restriction digest: by RE enzyme and amplification by PCR
2. Gel electrophoresis: SDS gel electrophoresis
3. Denaturation: Treating with HCl and NaOH
4. Blotting
5. Baking and Blocking with casein in BSA
6. Hybridization using labelled probes
7. Visualization by autoradiogram

Step I: Restriction digest

• The DNA is fragmentized by using suitable restriction enzyme. RE cuts the DNA at specific site generating fragments
• The number of fragments of DNA obtained by restriction digest is amplified by PCR

Step II: Gel electrophoresis

• The desired DNA fragments is separated by gel electrophoresis

Step III: Denaturation

• The SDS gel after electrophoresis is then soaked in alkali (NaOH) or acid (HCl) to denature the double stranded DNA fragments.
• DNA strands get separated

Step IV: Blotting

• The separated strands of DNA is then transferred to positively charged membrane nylon membrane (Nitrocellulose paper) by the process of blotting.

Step V: Baking and blocking

• After the DNA of interest bound on the membrane, it is baked on autoclave to fix in the membrane.
• The membrane is then treated with casein or Bovine serum albumin (BSA) which saturates all the binding site of membrane

Step VI: Hybridization with labelled probes

• The DNA bound to membrane is then treated with labelled probe
• The labelled probe contains the complementary sequences to the gene of interest
• The probe bind with complementary DNA on the membrane since all other non-specific binding site on the membrane has been blocked by BSA or casein.

Step VII: Visualization by Autoradiogram

• The membrane bound DNA labelled with probe can be visualized under autoradiogram which give pattern of bands.

Application of Southern blotting:

1. Southern blotting technique is used to detect DNA in given sample.
2. DNA finger printing is an example of southern blotting
3. Used for paternity testing, criminal identification, victim identification
4. To isolate and identify desire gene of interest.
5. Used in restriction fragment length polymorphism
6. To identify mutation or gene rearrangement in the sequence of DNA
7. Used in diagnosis of disease caused by genetic defects
8. Used to identify infectious agents

Southern Blotting: principle, procedure and application

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