Tools and Technologies Chapter Notes | Biology for BMAT (Section 2) PDF Download

Microscopy

• Microscopy is essential for biological studies, enabling visualization of structures too small for the naked eye.
• Advanced microscopy techniques allow highly magnified and three-dimensional imaging of minute structures, including bacterial and viral DNA molecules.
• The first microscope, designed by Robert Hooke in 1665, used magnifying lenses to observe cork slices, leading to the term "cell" for the honeycomb-like structure.
• Magnification, the ability to increase retinal image size, is calculated as the ratio of the retinal image size with a microscope to that without, or using the formula M = f / (f - d), where f is the focal length and d is the object distance.
• Compound microscopes, commonly used in laboratories, have two lens sets: an objective lens near the object and an eyepiece for viewing, with total magnification being the product of their magnifying powers (M₀ × Mₑ).
• Resolving power, the ability to distinguish two closely situated objects, is measured by the smallest distance between two points.
• A compound microscope consists of a base, stage with a central hole, arm, body tube, nosepiece with multiple objective lenses (4×, 10×, 40/45×, 100×), eyepiece (10× or 15×), adjustment screws, light source, and condenser for focusing light.
• Bright field microscopy uses light to illuminate stained objects, with common stains like carmine, eosin, safranin, methylene blue, and giemsa enhancing visibility.
• Dark field microscopy blocks central light, using oblique beams to illuminate objects against a dark background, ideal for detecting mitochondria, nuclei, and vacuoles.
• Phase contrast microscopy alters light wave amplitude and phase based on object density, enhancing contrast for studying cell organelles and chromosomes without staining.
• Fluorescence microscopy uses fluorophores (e.g., acridine orange, bisbenzimide) that emit longer-wavelength light when illuminated, aiding in identifying bacteria, viruses, or specific organelles for infection diagnosis.
• Electron microscopy employs electron beams with wavelengths 100,000 times shorter than visible light, magnified by electromagnetic lenses in a vacuum, viewed on a fluorescent screen.
• Transmission electron microscopy (TEM) passes electrons through ultra-thin, heavy metal-coated specimens to create high-resolution images.
• Scanning electron microscopy (SEM) uses reflected electrons from gold or platinum-coated surfaces to generate highly magnified, resolved surface images.
• Confocal microscopy, a sophisticated technique, resolves detailed structures in fluorescently labeled fixed cells or tissues, producing sharp, high-resolution images.

Centrifugation

• Centrifugation separates particles or molecules based on density under centrifugal force, achieved by spinning them in a solution at high speed around an axis.
• The centrifuge, the equipment used, includes a base, a rotating container (rotor), and a lid, with centrifuge tubes holding the cell extract or mixture.
• Spinning at a desired speed (revolutions per minute, rpm) for a specific time causes particulate material to settle at the tube bottom.
• Differential centrifugation separates particles based on sedimentation rates, influenced by size and density, used for isolating nuclei, mitochondria, chloroplasts, or large proteins.
• Density-gradient centrifugation separates particles of similar size but different densities using a density gradient in tubes, with heavier molecules sedimenting outward and lighter ones remaining inward.
• Ultracentrifugation, performed at speeds of 100,000×g or more, separates molecules with high precision, using table-top/clinical centrifuges, high-speed centrifuges, or ultracentrifuges.

Electrophoresis

• Electrophoresis separates macromolecules based on their charge-to-mass ratio under an electric field, derived from the Greek terms "electro" (electricity) and "phoresis" (migration).
• First observed in 1807 by Russian professors Peter Ivanovich Strakhov and Ferdinand Frederic Reuss, who noted clay particle migration in water under a constant electric field.
• Biological molecules like nucleotides, DNA, RNA, and proteins, bearing ionizable groups, exist as charged species (cations or anions) at a given pH, migrating toward the cathode or anode based on net charge.
• Molecule mobility is inversely proportional to size and directly proportional to charge, enabling separation.
• Agarose gel electrophoresis uses a matrix of agarose, a polysaccharide forming a solid, porous gel when heated in buffer and cooled, placed in a gel box with positive and negative electrodes.
• The gel box contains a salt-containing buffer to conduct current, with DNA-loaded wells positioned toward the negative electrode and DNA migrating toward the positive electrode (anode) due to its negative charge from phosphate groups.
• DNA fragments separate by size, with shorter fragments moving faster through gel pores, run at 80-120V, and visualized using a UV trans-illuminator with ethidium bromide (EtBr) staining, which intercalates DNA and fluoresces under UV light.
• EtBr, a potential carcinogen, requires careful handling and can be mixed into the gel, buffer, or applied post-run.
• Polyacrylamide gel electrophoresis (PAGE) separates proteins by size, using sodium dodecyl sulfate (SDS) to impart a uniform negative charge, with polyacrylamide gels formed by polymerizing acrylamide and N,N'-methylene-bis-acrylamide.
• Gel hardness, controlled by acrylamide concentration (4-8% for loose gels, 12-20% for hard gels), affects macromolecule friction, with loose gels allowing faster migration of high molecular weight molecules and hard gels favoring smaller molecules.
• Tracking dyes like bromophenol blue, xylene cyanol, or Orange G, added to samples, track migration progress, with bromophenol blue being highly mobile and stopping electrophoresis when it reaches the anode.
• Proteins are visualized using Coomassie Brilliant Blue R-250, fixed with methanol and acetic acid, and destained to reveal blue protein bands, with sizes determined by comparison to a molecular weight ladder.

Enzyme-Linked Immunosorbent Assay (ELISA)

• ELISA, invented by Eva Engvall and Peter Perlman in 1971, quantitatively measures antigen or antibody concentrations via enzyme-catalyzed reactions monitoring antigen-antibody interactions.
• An enzyme-conjugate linked to a specific antibody produces a colored product, measured by an ELISA reader or spectrophotometer, offering a safer, cost-effective alternative to other immunological assays.
• Direct ELISA involves coating microtiter wells with antigen, adding enzyme-conjugated antibody to bind directly, and measuring the colored product, being faster but prone to nonspecific binding.
• Indirect ELISA detects antibodies in two stages: unlabelled primary antibody binds to antigen-coated wells, followed by enzyme-conjugated secondary antibody binding to the primary, enhancing sensitivity but requiring washing to remove unbound antibodies.
• Sandwich ELISA uses a capture antibody coated on wells to bind antigen, followed by a second enzyme-conjugated antibody targeting a different epitope, offering high specificity without needing antigen purification.
• Competitive ELISA measures antigen by incubating antibody with antigen, adding the mixture to antigen-coated wells, and using enzyme-conjugated secondary antibody to detect bound primary antibody, with higher antigen concentrations reducing color intensity.
• ELISA applications include estimating antigen or antibody presence, determining serum antibody concentrations for virus tests, detecting food allergens, and tracking disease outbreaks (e.g., HIV, bird flu, cholera).

Chromatography

• Chromatography separates mixture components using a mobile phase (solution with solutes and eluent) and a stationary phase (adsorbent, ion-exchange resin, porous solid, or gel).
• Separation relies on solutes’ differential migration rates based on their affinity for the stationary phase, packed in a cylindrical column, producing a chromatogram of solute peaks.
• Liquid chromatography serves as both an analytical and preparative technique for biomolecule purification, particularly for high-purity therapeutics and pharmaceuticals.
• Adsorption chromatography (ADC) uses polar adsorbents (e.g., silica gel, alumina) to adsorb solute molecules, with solvent polarity influencing separation.
• Liquid-liquid partition chromatography (LLC) separates solutes based on their partition coefficients between two immiscible solvents, with one fixed as the stationary phase.
• Ion-exchange chromatography (IEC) adsorbs solute ions onto ion-exchange resin via electrostatic forces, used for fractionating antibiotics and proteins, with elution adjusted by pH or ionic strength.
• Gel filtration chromatography, also known as size exclusion or molecular sieve chromatography, separates molecules by size using porous gel particles (e.g., cross-linked dextrans, agaroses, polyacrylamide), suitable for proteins and lipophilic compounds.
• Affinity chromatography (AFC) exploits biomolecule binding specificity, using ligands linked to the column to retain solutes like enzymes, antibodies, or nucleic acids with high selectivity.
• High-performance liquid chromatography (HPLC) uses small stationary phase particles, requiring high-pressure pumping due to flow resistance, enhancing resolution.
• Gas chromatography (GC) separates volatile components (e.g., alcohols, ketones) using a gas mobile phase, widely used but less relevant to bioprocessing compared to liquid chromatography.

Spectroscopy

• Spectroscopy studies the interaction of electromagnetic radiation with matter, used for identifying substances, estimating colored compound concentrations, analyzing chemical structures, and studying intermolecular bonding.
• A spectrophotometer includes a light source, collimator, monochromator, and wavelength selector to transmit specific wavelengths for analysis.
• Colorimetry measures light transmittance and absorbance in colored solutions, using a tungsten-filament lamp, colored filters, cuvette, and photocell, based on the Beer-Lambert law (A = εlc), where absorbance (A) is proportional to concentration (c) and path length (l).
• UV-visible spectrophotometry measures light absorption in UV and visible regions using a prism or grating for monochromatic light, with components like light sources, monochromator, cuvette, detector, and signal processor.
• Spectroscopic techniques include UV/visible spectroscopy (detects functional groups), atomic absorption spectroscopy (measures metal levels), infrared spectroscopy (identifies functional groups), Raman spectroscopy (identifies contaminants), nuclear magnetic resonance (NMR) spectroscopy (provides structural information), and fluorescence/phosphorescence spectroscopy (detects organic compounds).

Mass Spectrometry

• Mass spectrometry identifies unknown compounds, quantifies materials, and elucidates molecular structures by converting samples into gaseous ions, characterized by mass-to-charge ratios (m/z) and relative abundances.
• The mass unit is Dalton (Da), defined as 1/12th the mass of a carbon-12 atom.
• Components include an ion source (produces gaseous ions), an analyzer (resolves ions by m/z), and a detector (records ion abundance).

Fluorescence In Situ Hybridisation (FISH)

• FISH, a cytogenetic technique, uses fluorescent molecules to bind complementary chromosome regions, identifying gene locations and chromosomal abnormalities.
• It involves a fluorescent DNA probe and a chromosome target sequence, observed under a fluorescence microscope.
• The process includes designing a fluorescently labeled probe (e.g., with fluorescein), denaturing chromosomes and the probe, hybridizing the probe to complementary sites, washing off excess probe, and observing fluorescent signals.
• Applications include chromosome painting, where multifluor probes label each chromosome with unique colors, visualized under a fluorescence microscope to generate karyotypes.

DNA Sequencing

• DNA sequencing determines the order of nucleotides (A, T, G, C) in DNA, critical for understanding genetic information and developmental programs.
• Sequence alterations can lead to non-functional proteins, affecting health, disease susceptibility, and drug responses.
• Sequencing is vital for studying gene structure, function, evolution, and applications in diagnostics, forensics, agriculture, and environmental research.
• Sanger’s method (enzymatic, dideoxynucleotide chain termination) uses dideoxynucleotides (ddNTPs) lacking a 3’ -OH group, terminating DNA synthesis when incorporated.
• In Sanger’s method, template DNA is mixed with a primer, labeled dNTPs, and split into four tubes (A, C, G, T), each spiked with a different ddNTP, with DNA polymerase synthesizing strands until termination.
• Fragments are separated by high-resolution PAGE, visualized via radioactive or fluorescent labels, and read from the gel bottom upward.
• Maxam and Gilbert’s method (chemical degradation) labels DNA at the 5’ end, cleaves it base-specifically with chemicals (e.g., dimethylsulphate for G, hydrazine for C+T), and separates fragments by PAGE for sequence determination.
• Automated sequencing, based on Sanger’s method, uses fluorescently labeled ddNTPs (e.g., Rhodamine 110 for A, Rhodamine 6G for C), capillary tubes, and a single-tube reaction, with a laser and fluorimeter detecting fluorescence for rapid, cost-effective sequencing.
• Next-generation sequencing (NGS) platforms (e.g., Roche/454, Solexa/Illumina, SOLiD) enable massively parallel sequencing for whole genomes, significantly faster than traditional methods.

DNA Microarray

• DNA microarray technology, a high-throughput hybridization-based method, analyzes thousands of DNA fragments simultaneously to quantify gene expression.
• It relies on complementary DNA strand pairing via hydrogen bonds, with single-stranded DNA (ssDNA) probes immobilized on glass or nylon chips.
• Probes, arranged in rows and columns, are cDNAs, PCR amplicons, or oligonucleotides corresponding to mRNAs, with oligonucleotide probes being popular for analyzing thousands of genes.
• A typical experiment involves mRNA extraction, probe labeling, hybridization and washing, and scanning/data analysis.
• mRNA, extracted to quantify gene activity, is converted to stable cDNA via reverse transcription.
• cDNA fragments are labeled with fluorochromes (e.g., Cy3, Cy5), hybridized to chip probes to form duplexes, and unbound fragments are washed away.
• Chips are scanned with sophisticated scanners, and software analyzes fluorescence intensity to quantify gene expression, with green indicating upregulated genes and red indicating downregulated genes.

Flow Cytometry

• Flow cytometry analyzes cells’ physical or chemical properties for qualitative or quantitative measurements, tracing back to the 1950s Coulter counter, which measured particle volume via impedance changes.
• Modern flow cytometry passes cells one by one through a fluid stream, with laser beams detecting properties or fluorescent labels, and sensors capturing deflected light.
• Detectors, aligned with or perpendicular to the laser, assess surface or internal properties, generating comprehensive cell images.
• Fluorescently labeled antibodies identify antigens or proteins, enabling localization in cells, such as immune system cells.
• Cells labeled with different fluorescent dyes can be mechanically separated by loading them onto a charging electrode, releasing them drop-wise, and detecting fluorescence with laser-based sensors, recorded as data plots.