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1. MOLECULAR METHODS OF
DIAGNOSIS
DIVYA RAVALI B

2. MOLECULAR METHODS
 PCR
 DNA SEQUENCING
 RT PCR
 WESTERN BLOT
 FLUORESCENCE IN-SITU HYBRIDIZATION
 NUCLEIC ACID ARRAYS
 PROTEOMICS WITH MASS SPECTROMETRY
 TRANSGENIC MICE

3. POLYMERASE CHAIN REACTION
Purpose
• Amplify a specific piece of DNA from a complex mixture
Requirements
• Need to know sequence of the DNA of interest (at least of its ends)
Outline of method
• Oligonucleotide primers are designed to hybridize to specific sequences at each end of the DNA of interest.
These primers are added to a reaction vessel mixture containing the template DNA along with a
thermostable DNA polymerase, the nucleotides dATP (A), dTTP (T), dGTP (G) and dCTP (C), and buffer
• The reaction vessel is placed in a thermal cycler, which controls the temperature of the reaction through
many cycles
• Each cycle contains the following steps: (1) denaturation; (2) primer annealing or primer hybridization; (3)
primer extension; and (4) repeat of the complete cycle of PCR 30–40 times

5. Benefits
• simple and rapid
• Because the PCR product is exponentially increased, it is extremely sensitive in amplifying low amounts
of DNA. Each cycle increases the number of PCR products twofold. The total number of PCR products
after n cycles will be 2

6. Experimental applications
• DNA can be amplified either for detection of a specific sequence or for cloning that
sequence
• PCR can be used to label DNA with fluorescent or radioactive nucleotides
• PCR can be used for rapid haplotype analysis
Modifications/alternatives
• Quantitative PCR
• Southern blot, in situ hybridization, comparative genomic hybridization

7. DNA SEQUENCING
Purpose
• Determine the sequence or order of nucleotides (A, G, C, T) in a stretch of DNA
Requirements
• The piece of DNA to be sequenced can be either a PCR product or a cloned piece of DNA present in
a plasmid, but it should be pure

8. Underlying concepts
• Chain termination (Sanger sequencing), or a variation thereof, is a standard method for
DNA sequencing
• In chain termination, the extension of a new strand of DNA is stopped by the addition of an
analogue of dATP, dCTP, dGTP or dTTP (ddATP, ddCTP, ddGTP or ddTTP, respectively) to
the sequencing mixture. When the DNA polymerase incorporates the analogue nucleotide
instead of the correct normal nucleotide, DNA synthesis is terminated because the
polymerase is no longer able to link to the next nucleotide
• Gel electrophoresis is used to separate the different sizes of DNA fragments that result from
chain termination synthesis. The DNA fragments are forced to travel through a gel using an
electric current;
the smaller molecules are less impeded by the gel and travel faster than larger molecules

9. Outline of method
• An oligonucleotide primer hybridizes to the DNA to be sequenced and DNA polymerase synthesizes a
second complementary strand
• The synthesis of the second strand is interrupted randomly by the incorporation of the fluorescent
nucleotide analogues (ddATP, ddGTP, ddCTP, ddTTP), and the DNA fragments containing this final
nucleotide analogue can be identified because each of the four ddNTPs is labeled with a different color
fluorochrome
• The different DNA fragments are electrophoresed through a polyacrylamide gel or capillary tubes
• The different-length DNA strands terminating with different fluorochrome-labeled nucleotide analogues
pass a fluorescence detector and indicate the order of the DNA sequence

11. Benefits
• The fluorescent chain termination method is able to rapidly sequence large amounts of
DNA with automated analysis of results
Experimental applications
• Determine previously unknown sequence
• Confirm sequence following the cloning of a DNA fragment of interest and other
manipulations
Modifications/alternatives
• Pyrosequencing
• Next-generation sequencing

12. REVERSE TRANSCRIPTION PCR
Purpose
• To amplify mRNA by PCR, the mRNA is first converted to DNA (called complementary DNA or
cDNA), followed by PCR amplification of a specific region of the cDNA to detectable levels
Requirements
• Starting material can be total cellular RNA (including ribosomal, transfer and messenger RNA
[mRNA]) or purified mRNA
Underlying concept
• In order to facilitate studies of RNA, many techniques that study RNA first convert the RNA to
cDNA with an enzyme called reverse transcriptase, an RNA-dependent DNA polymerase

13. Outline of method
• Reverse transcriptase can convert mRNA to cDNA by three different methods, depending on the primer
used for the initial RT step
(1) Random hexamer primers contain six nucleotides (6-mer) that have all possible sequence
combinations of the dA, dG, dC and dT nucleotides (46 possible combinations). These random
hexamers will hybridize to the corresponding complementary sequences in the sample RNA.
(2) Oligo dT primers contain only dT nucleotides and hybridize to the complementary string of dA
nucleotides that are present at the end of mRNA molecules (poly A tail)
(3) The third choice is a primer that will only hybridize to a specific mRNA sequence
• After the mRNA has been converted to cDNA, primers that can hybridize to specific sequences are added
and PCR amplification is performed

15. Benefits
• As for PCR, RT-PCR is simple and rapid
• RT-PCR is extremely sensitive in detecting low levels of mRNA transcripts
Experimental applications
• The mRNA gene transcripts can be amplified for subsequent cloning or sequencing
• The mRNA gene transcripts (instead of the gene) could be analyzed for
the presence of mutations

16. Modifications/alternatives
• Quantitative RT-PCR
• Alternative methods to measure RNA levels include digital PCR, in
which the sample is partitioned into individual nucleic acids, and
Northern blots

17. WESTERN BLOT
Purpose
• Western analysis can measure the size and the amount of protein present in a sample
Requirements
• Western analysis requires an antibody that is specific for the protein of interest (i.e. does
not cross react with other proteins

18. Outline of method
• A solubilized protein mix is separated on a
polyacrylamide gel and transferred electrophoretically
to a membrane. The membrane is then soaked in a
buffer containing the antibody. Bound antibody is
detected by a chromogenic or
chemiluminescent assay

19. Benefits
• Western analysis is a simple and sensitive method to detect and quantify
proteins present in a complex mixture
• Western analysis can determine the molecular weight of a specific protein
relative to standard controls
Experimental applications
• Detection, quantification, and characterization of a specific protein
• Identification of antibody activity to a known antigen

20. Modifications/alternatives
• Dot blot – a drop of the protein mixture is placed on a paper membrane and the protein of interest
detected with antibodies, as in the Western blot.
• Immunoprecipitation (IP) – the specific antibody is added to the protein mixture and the resulting
antibody–protein complexes are then isolated.
• IP–Western – protein–protein interactions can be studied by first immunoprecipitating the protein with
one antibody, bringing down a protein complex. The protein complex is then separated on a
polyacrylamide gel, followed by Western blotting to detect members of the protein complex

21. • Enzyme-linked immunosorbent assay (ELISA) – this is a sensitive and specific method for quantifying
the amount of a protein. The protein of interest is captured on a plate coated with a monoclonal antibody
and other proteins are washed away. The protein is then detected using a second antibody that has been
modified for detection using a colorimetric or luminescent assay
• Immunohistochemistry – this is used to visualize the cellular localization of a protein. An antibody to the
protein of interest is applied to a tissue section. The antibody is detected using a secondary antibody
coupled to an enzyme that reacts with a substrate to produce a colored precipitate

22. FLUORESCENCE IN SITU
HYBRIDIZATION (FISH)
Purpose
• To visualize locations and levels of specific sequences of DNA or RNA in tissue sections
Requirements
• Tissue section of formalin-fixed paraffin-embedded tissue or fresh-frozen tissue
• The target sequence must be known
Underlying concepts
• A fluorescently labeled nucleic-acid probe hybridizes to complementary sequences in tissue
sections
• The signal is detected using fluorescence microscopy

23. Outline of method
• A probe is designed that is complementary to the target sequence and is directly or
indirectly labeled with a fluorochrome
• Nucleic acids in the tissue section are denatured and then the FISH probe is applied to
the tissue section to hybridize to the target sequence
• Unbound probe is washed away
• Samples are viewed by fluorescence microscope

25. Benefits
• Able to identify which cells exhibit a specific abnormality in DNA, such as gene deletion, amplification,
or translocation
• Able to determine which cells are expressing a particular gene
Experimental applications
• Identification of DNA copy number aberrations in cancer cells
• Determining the spatial pattern of gene expression in a tissue

26. Modifications/alternatives
• Single molecule RNA FISH
• FISH of metaphase preparations to diagnose chromosomal abnormalities
• Spectral karyotyping or multifluor FISH to detect chromosomal translocations and large
deletions or duplications
• PCR or RT-PCR following laser microdissection of pure cell population
• Array-based comparative genomic hybridization (CGH)

27. NUCLEIC ACID ARRAYS
Purpose
• To profile the mRNA expression of thousands of genes in one experiment
Requirements
• Total RNA or mRNA from samples (larger amounts than required for RT-PCR)
Underlying concepts
• Hybridization to DNA – the same principle of hybridization applies as in PCR, except that
many different genes are being
evaluated simultaneously. DNA is attached to beads, chemically synthesized on a surface
at thousands of specific locations, or spotted onto glass slides. If cells are expressing
mRNA of the corresponding gene, labeled cRNA prepared from mRNA in those cells will
hybridize and generate a signal
intensity related to its level of expression

28. Outline of method

29. Benefits
• Thousands of genes can be rapidly and quantitatively profiled in one experiment
Limitations/errors
• Genes expressed at low levels may not be detected
• For some genes, the cRNA may not hybridize under the conditions used
• The RNA must be of very high quality
Experimental applications
• Microarray analysis can be used to profile changes in gene expression
• The patterns of gene expression can be used to group samples

30. Modifications/alternatives
• RNA-seq
• MicroRNA arrays
• Comparative genomic hybridization (CGH) to detect differences in DNA copy
number
• Single nucleotide polymorphism (SNP) arrays to assess markers of genetic
variation; useful for linkage analysis, including genome-wide
association studies

31. PROTEOMICS WITH MASS
SPECTROMETRY
Purpose
• High-throughput analysis that allows investigators to rapidly and quantitatively assess the complex
mixtures of proteins present in a cell at a given point in time
Requirements
• A mixture of cellular proteins, or peptides derived from these proteins, is purified from a defined
population of cells and then separated into proteins and/or peptides prior to mass spectrometry
Underlying concepts
• Mass spectrometry is able to measure very precisely the size or mass of proteins, peptides, or peptide
fragments by giving these peptides a positive charge (ionization) and then measuring the time required
for the positively charged peptide ions to move through a tube to a detector
(time-of-flight)

32. Outline of method

33. Benefits
• Mass spectrometry is extremely sensitive and able to detect very small amounts of proteins/peptides
Experimental applications
• Mass spectrometry can be used to determine the complete set of proteins present in a defined
population of cells
• Because of the sensitivity of mass spectrometry analysis, it holds promise as a sensitive diagnostic tool

34. Modifications/alternatives
• Isotope labeling approaches for quantitative mass spectrometry, including isotope-coded
affinity tag (ICAT), stable isotope labeling by amino acids in cell culture (SILAC), tandem mass tags
(TT), and isobaric tags for relative and absolute quantification (iTRAQ)
• Label-free quantitative proteomics
• Antibody protein arrays and reverse-phase protein arrays

35. TRANSGENIC MICE
Purpose
• Transgenic mouse models allow investigators to study the effects of a transgene (the gene of interest) on
a cellular, tissue, and whole animal level. The transgene can be selectively expressed in a particular cell
type or tissue by using a promoter/enhancer regulatory region that is specific for that cell type
Requirements
• A transgene or gene of interest, whose biologic function is to be characterized in an animal model
• A regulatory region (promoter/enhancer) that will selectively express the transgene in a specific tissue
• The facilities and technologies to create transgenic mice

36. Outline of method
• A transgene construct, defined as the transgene and a regulatory region (promoter/enhancer), is
prepared for injection
• The transgene is microinjected into fertilized eggs (single-cell stage) and the transgene integrates into
the genome, usually at a single site
• These injected eggs are then implanted into a recipient mother, who then gives birth to a
heterozygous “founder” mouse that can be either male or female.
The founder mouse is referred to as a transgenic mouse because it only contains the transgene on one
of the two paired chromosomes (e.g. only one
copy of chromosome 7 contains the transgene, whereas the other chromosome 7 does not)

37. • The founder mice are bred with normal non-transgenic mice of the same strain. The progeny mice
will be both heterozygous transgenic and non-transgenic, according to Mendelian genetics
• In order to derive homozygous transgenic mice that contain the transgene on both of the paired
chromosomes (e.g. both copies of chromosome 7 contain the transgene), two heterozygous
transgenic mice are mated. Homozygous transgenic mice will have a double dose of the transgene,
which may lead to different phenotypes and biologic effects from those present in the heterozygous
transgenic mice

39. Benefits
• Transgenic mouse models can help us understand the in vivo biologic effects of
known and unknown genes when expressed in a specific tissue
Experimental applications
• Transgenic mouse models of disease can be used to test the effectiveness of new
therapeutic agents

40. Modifications/alternatives
• More precise control of transgene expression can be achieved with promoters/enhancers
that are inducible and can be regulated (i.e. turned off and on at different time points)
• Programmable nuclease systems for RNA-guided genome editing such as transcription
activator-like effector nuclease (TALEN) and clustered regularly
interspaced short palindromic repeat (CRISPR)/CRISPR-associated protein (Cas)
• “Knockout” transgenic mouse models allow investigators to understand the biologic effects
of gene mutations and deletions that cause a loss of specific proteins

41. KNOCKOUT TRANSGENIC MICE
Purpose
• Knockout transgenic mice have both alleles of a normal endogenous gene modified, mutated,
or deleted in all cells (whole animal or targeted tissue) so that protein can no longer be
produced by that gene. Therefore, knockout transgenic mice allow investigators to understand
what happens to a cell or tissue when a particular gene can no longer be expressed
Requirements
• Murine embryonic stem cells (ES cells) that are totipotent and capable of forming and
reconstituting all tissues and organs of a mouse
• A targeting construct defined as a defective gene that can hybridize to the normal endogenous
gene in ES cells and modify or delete the targeted gene so that it is no longer able to express a
functional protein

42. Outline of method
• A targeting vector that contains some sequences of the gene to be targeted is created and
introduced into ES cells. This targeting vector is able to selectively hybridize to one of the
endogenous alleles of a gene, and through a process known as homologous recombination, the
targeting vector modifies or deletes the endogenous gene so that no normal protein is produced.
ES cells that contain this defective endogenous gene can be selected for and isolated
• The ES cells containing the “knocked out” gene are introduced into early mouse embryos,
known as blastocysts, which are then implanted into recipient mothers. The progeny will be
chimeric mice; all tissues and organs, including testes and ovaries, will contain a mixture of
normal cells and cells that lack the “knockout” gene allele

43. • The chimeric mice are next mated to normal mice. Depending on the degree of chimerism, a certain
percentage of the offspring will be heterozygous, containing one normal allele and one allele that has
been modified or deleted (knocked out). As described for the mating procedure in transgenic mice, the
genotypes of all progeny will follow the rules of Mendelian genetics
• To obtain completely “knocked out” mice, two heterozygous mice are mated. Approximately 25% of the
offspring will lack a normal endogenous gene in every cell and tissue

44. Benefits
• This approach yields mice that completely lack the gene product in all cells of the whole animal or
targeted tissues
Experimental applications
• Unlike regular transgenic mice, knockout transgenic mice allow investigators to study the biologic
effects of losing gene expression. This is particularly
useful in the study of genes that may normally suppress cancer development (tumor suppressors); the
tumor suppressor gene can be knocked out in all
mouse cells in vivo, and the mechanisms of cancer development in different tissues can be studied